National Wildlife Health Center

Anisakiosis and Pseudoterranovosis

Circular 1393

U.S. Department of the Interior

U.S. Geological Survey
1 2

3 4

5

6

7 8

Cover: 1. Common seal, by Andreas Trepte, CC BY-SA 2.5; 2. Herring catch, National Oceanic and Atmospheric Administration; 

3. Larval Pseudoterranova sp. in muscle of an American plaice, by Dr. Lena Measures; 

4. Salmon sashimi, by Blu3d, Lilyu, GFDL CC BY-SA3.0; 5. Beluga whale, by Jofre Ferrer, CC BY-NC-ND 2.0; 

6. Dolphin, National Aeronautics and Space Administration; 

7. Squid, National Oceanic and Atmospheric Administration; 8. Krill, by Oystein Paulsen, CC BY-SA 3.0.
Anisakiosis and Pseudoterranovosis

By Lena N. Measures

Edited by Rachel C. Abbott and Charles van Riper, III

USGS National Wildlife Health Center

Circular 1393

U.S. Department of the Interior

U.S. Geological Survey
ii

U.S. Department of the Interior

SALLY JEWELL, Secretary

U.S. Geological Survey

Suzette M. Kimball, Acting Director

U.S. Geological Survey, Reston, Virginia: 2014

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Suggested citation:

Measures, L.N., 2014, Anisakiosis and pseudoterranovosis: Reston, Va., U.S. Geological Survey Circular 1393, 34 p., 

2 appendixes, http://dx.doi.org/10.3133/cir1393.

Library of Congress Cataloging-in-Publication Data

Measures, Lena N., 1950- , author.

 Anisakiosis and pseudoterranovosis / by Lena N. Measures.

 p. ; cm. -- (Circular ; 1393)

 At head of title: National Wildlife Health Center.

 Includes bibliographical references and index.

 ISBN 978-1-4113-3809-8 (alk. paper)

 I. National Wildlife Health Center (U.S. : 1993-) II. Title. III. Series: U.S. Geological Survey circular ; 1393.

 [DNLM: 1. Anisakiasis--epidemiology. 2. Ascaridida Infections--epidemiology. WC 870]

 RC840.A56

 616.9’654--dc23

 2014032432

ISSN 1067-084X (print)

ISSN 2330-5703 (online)
iii

Foreword

C. van Riper, III, R. C. Abbott, C. Bunck, and M. Friend 

“Make no mistake, they are connected, these disease outbreaks coming one after 

another. And they are not simply happening to us; they represent the unintended 

results of things we are doing. They reflect the convergence of two forms of crisis on 

our planet. The first crisis is ecological, the second is medical.” 

David Quammen, Spillover

The emergence of many new (and older) zoonotic diseases in humans in recent years is largely 

a result of our densely populated, highly mobilized, and environmentally disrupted world that 

is experiencing rapid climatic changes. The number of zoonotic diseases detected has been of 

unprecedented scope relative to geographic areas of occurrence, wildlife species affected, and 

the variety of pathogens involved (Daszak and others, 2000; Friend, 2006). As towns and cities 

expand, the wildland-urban interface broadens and human associations with wildlife become 

increasingly frequent. With geographic distance and isolation no longer serving as meaningful 

barriers, opportunities for once isolated diseases to spread have never been greater. Consequently, and especially as a result of rapid warming in the polar region due to climate change, 

zoonotic diseases such as anisakiosis and pseudoterranovosis will begin to appear more frequently in northern latitude animal and human populations.

Human infections with “whale worm” or “seal worm” are becoming more common with the 

popularity of eating raw fish. People are accidentally infected by larval stages of the nematodes 

in marine fish or squid that are raw, poorly cooked, cold-smoked, lightly salted, or marinated. 

It is safe to assume that both these parasites can be found wherever suitable hosts occur and 

favorable environmental conditions permit. Thus, humans consuming improperly prepared 

infected wild fish or squid are at risk, especially those with cultural culinary practices that do 

not kill larvae. And as gastronomic trends spread around the globe, foodborne zoonoses, such as 

anisakiosis and pseudoterranovosis, will occur in new locations.

Dealing with emerging zoonotic diseases requires the ability to recognize pathogens when they 

first appear and to act quickly and appropriately. Because outbreaks often are evident in nonhuman components of the environment before humans are affected, understanding our environment and associated ‘sentinel’ wildlife is a prerequisite to protecting human health. Through 

understanding infection levels in wildlife populations, we will be better able to predict future 

human infection levels. This report, seventh in the series of U.S. Geological Survey Circulars on 

zoonotic diseases, will help us to better understand the routes of anisakiosis and pseudoterranovosis infections and how best to adequately monitor these zoonotic diseases.

Daszak, P., Cunningham, A.A., and Hyatt, A.D., 2000, Emerging infectious diseases of wildlife—Threats to biodiversity and human health: Science, v. 287, p. 443–449.

Friend, M., 2006, Disease emergence and reemergence: The wildlife-human connection: Reston, Va., U.S. Geological Survey Circular 1285, 388 p.

Quammen, D., 2012, Spillover: animal infections and the next human pandemic: New York, 

N.Y., W.W. Norton and Co., Inc., p. 39.

v

Contents

Foreword ...................................................................................................................................................... iii

Overview........................................................................................................................................................ 1

Background.....................................................................................................................................................1

Causative Agent.............................................................................................................................................3

Geographic Distribution................................................................................................................................6

Patterns and Trends ......................................................................................................................................6

Species Susceptibility...................................................................................................................................6

Human Infections..................................................................................................................................6

Marine Mammal Infections.................................................................................................................8

Fish and Invertebrates .........................................................................................................................8

Birds......................................................................................................................................................11

Other Animals .....................................................................................................................................11

Obtaining a Diagnosis .................................................................................................................................11

Disease Ecology...........................................................................................................................................13

Points to Ponder...........................................................................................................................................18

Disease Prevention and Control................................................................................................................18

References Cited .........................................................................................................................................20

Glossary.........................................................................................................................................................26

Appendixes ..................................................................................................................................................29

Topic Highlight Boxes 

Culture, Technology, and Anisakiosis ........................................................................................................2

A Case of Human Anisakiosis......................................................................................................................7

Migration of Adult Hysterothylacium aduncum in Atlantic Cod After Host Death ...........................16

Commercial Candling of Fish Fillets to Visualize and Extract Pseudoterranova decipiens ............17

The Baermann Technique ..........................................................................................................................19

Figures

1. Photo showing adult Anisakis simplex in the stomach of a pilot whale .......................................9

2. Photo showing adult Pseudoterranova decipiens in the stomach of a grey seal.......................9

3. Photo showing red vent syndrome in Atlantic salmon ..................................................................10

4. Anatomical drawing of key morphological features of anisakid larvae removed from 

fish: A, Anisakis; B, Contracaecum/Phocascaris; C, Pseudoterranova................................12

5. Diagram showing the life cycle of Anisakis simplex......................................................................13

6. Photo showing larval Anisakis sp. (arrows) in the abdominal cavity of an 

Atlantic herring...............................................................................................................................14

7. Diagram showing the life cycle of Pseudoterranova decipiens ..................................................15

8. Photo showing larval Pseudoterranova sp. in muscle of an American plaice ..........................15
vi

Tables

1. Taxonomic classification of the parasites causing anisakiosis and

pseudoterranovosis .........................................................................................................................1

2. Timeline of events in the discovery of human anisakidosis............................................................2

3. Marine mammal hosts and geographic range of Anisakis spp. ....................................................4

4. Marine mammal hosts and geographic range of Pseudoterranova spp. ....................................5

5. Forms of human anisakidosis...............................................................................................................7

6. Methods to reduce risk of infection by anisakid larvae for consumers of fish .........................18

Conversion Factors and Abbreviations

International System of Units to Inch/pound

Multiply By To obtain

Length

centimeter (cm) 0.3937 inch (in.)

millimeter (mm) 0.03937 inch (in.)

Mass

kilogram (kg) 2.205 pound avoirdupois (lb)

Temperature in degrees Celsius (°C) may be converted to degrees 

Fahrenheit (°F) as follows:

°F=(1.8×°C)+32

Words shown in bold are defined in the glossary.
Anisakiosis and Pseudoterranovosis

By Lena N. Measures1

Synonyms 

Anisakiasis, whaleworm and herringworm infection, 

pseudoterranovosis, sealworm, and codworm infection

Overview

Anisakiosis is a parasitic disease caused by infection 

with larval nematodes or roundworms of the taxonomic

genus Anisakis, which belong to the family, Anisakidae, subfamily Anisakinae. Anisakiosis is a zoonotic disease, meaning 

it is transmissible between animals and humans and vice versa. 

Other related roundworms, such as Pseudoterranova and Contracaecum, may cause similar, but less frequently reported, 

diseases (pseudoterranovosis, contracaecosis). The terms 

anisakidosis (anisakidiasis), referring to disease caused by 

members of the family, and anisakinosis (anisakiniasis), referring to disease caused by members of the subfamily, may be 

encountered in technical literature (Kassai and others, 1988) 

when specimens cannot be identified to genus. Because most 

cases of anisakidosis in humans are due to larval Anisakis spp. 

and, to a lesser extent, larval Pseudoterranova spp., further 

discussion will be limited to these anisakids (table 1).

The life cycles of Anisakis spp., commonly called whaleworm (previously known as herringworm), and Pseudoterranova spp., commonly called sealworm (previously known 

as codworm), are complex and involve three marine hosts

(invertebrates, fish, and marine mammals). Whales, dolphins, 

or porpoises (collectively known as cetaceans) are the definitive hosts in which Anisakis spp. become sexually mature, 

and seals, sea lions, or walrus (collectively known as pinnipeds) are the definitive hosts of Pseudoterranova spp. These 

zoonotic parasites have medical and economic importance and 

can result in considerable costs to the fishing industry. Humans 

are accidentally infected by consuming raw, poorly cooked, 

cold smoked, lightly salted, or marinated marine fish or squid, 

the intermediate hosts infected with larval stages. Human 

infections are becoming more common with the popularity of 

eating raw fish as well as improved medical diagnostics. Infections are rarely diagnosed correctly in North America, which 

often leads to surgical intervention. Symptoms in humans may 

involve nausea, vomiting, and severe stomach (gastric) or 

abdominal pain.

1

 Maurice Lamontagne Institute, Fisheries and Oceans Canada.

Table 1. Taxonomic classification of the parasites causing 

anisakiosis and pseudoterranovosis. 

Classification Designation

Kingdom Animalia

Phylum Nematoda

Class Secernentea

Order Ascardida

Family Anisakidae

Subfamily Anisakinae

Genus Anisakis Pseudoterranova

Species simplex 

sensu stricto

decipiens 

sensu stricto

berlandi krabbei

pegreffi bulbosa

typica azarasi

physeteris cattani

ziphidarum decipiens E

paggiae kogiae

brevispiculata ceticola

nascettii

schupakovi

Background

Although the genus of Anisakis was established in 1845 

by Félix Dujardin to group some parasitic nematodes known 

from the stomach and intestine of marine mammals, it took 

more than 100 years until the first human infections were 

described in Holland and Japan (table 2). Since then, thousands of people in Japan have contracted anisakiosis.

In the Western hemisphere, infections caused by larvae 

of the Anisakidae family or similar larvae were reported from 

Inuit in Greenland as early as 1867 (Martin, 1921 cited in 

Jackson, 1975) and Alaska (Hitchcock, 1950). Since then 

many human cases of “anisakiasis” have been reported from 

Canada and the United States, but few cases of anisakiosis 

have been confirmed by identification of specimens as Anisakis spp. (Smith, 1999; Couture and others, 2003). Most identified specimens from human cases in the Western Hemisphere 

are reported as Pseudoterranova spp., thus infections are 

called pseudoterranovosis (Amin and others, 2000).
2 Anisakiosis and Pseudoterranovosis

Table 2. Timeline of events in the discovery of human anisakidosis.

Date Event

1845 Genus of Anisakis is established by Félix Dujardin to 

group nematodes causing lesions in the stomachs 

of cetaceans.

1867 Ascarid-like or anisakine infections are reported from 

Inuit people in Greenland (Martin, 1921 cited in 

Jackson, 1975).

Early 

1940s

Gastrointestinal lesions in humans, with small 

Ascaris-like larvae in a few cases are documented 

in Japanese medical journals (Oshima, 1972).

1949 Ascarid-like or anisakine infections are reported from 

Inuit people in Alaska (Hitchcock, 1950).

1955 The first human cases of anisakiosis are observed in 

Holland (van Thiel and others, 1960; Rodenburg 

and Wielinga, 1960; Kuipers and others, 1960).

1955–67 Almost 150 cases of human anisakiosis are documented 

in Holland.

1965 The first human case of anisakiosis in Japan is 

described (Asami and others, 1965).

Pseudoterranovosis has been reported in Japan, but it 

is less common than anisakiosis (Ishikura and others, 1992). 

Three hundred thirty-five human cases of pseudoterranovosis 

in Japan and 19 in other countries, including the United States, 

Canada, Greenland, Chile, the United Kingdom, and Korea, 

have been reported (Ishikura and others, 1992). Pseudoterranovosis has been reported frequently in northern Japan 

(Hokkaido) and attributed to dietary differences compared to 

southern Japan and the presence of populations of pinnipeds, 

such as sea lions and seals. Recently, pseudoterranovosis has 

been reported from Mexico and South America (Torres and 

others, 2007).

For further reading, the bibliographic citations of key 

publications that address various aspects of anisakiosis and 

pseudoterranovosis are listed in appendix 2.

Box Title Here

Culture, Technology, and Anisakiosis

Anisakiosis in a human was first observed in 1955 in Holland (van 

Thiel and others, 1960; Rodenburg and Wielinga, 1960; Kuipers 

and others, 1960). Although larval forms of roundworms in the Anisakidae family are common in marine fish, the apparent sudden 

appearance of anisakiosis in Holland was attributed to changes 

in fish processing and marketing practices during the early 1950s. 

Traditionally, fresh-caught herring were eviscerated at sea, but 

with the advent of refrigeration, onboard ship processing changed 

to icing whole fish and eviscerating them when ships returned to 

port, sometimes days later. In addition, so called “green herring,” 

which was lightly salted raw herring, was introduced to the Dutch 

market and became very popular. These two practices led to larval 

Anisakis spp. migrating after the death of the fish host from the 

viscera into the muscles, surviving further inadequate processing 

and remaining alive for days. Consequently, almost 150 cases of 

human anisakiosis were documented between 1955 and 1967 in 

Holland. Dutch parasitologists eventually linked larval nematodes from herring with cases of anisakiosis in humans and finally 

identified them as Anisakis spp. As a result of these findings, in 

1968 Dutch legislation required that fish be frozen within 12 hours 

of catch at –20 °C and kept frozen for 24 hours before further 

processing (Oshima, 1972). Although cases of anisakiosis fell in 

Holland after these measures were implemented, anisakiosis still 

affects fish consumers in Europe. Consumption of gravlax (cured 

salmon, often dried) in Nordic countries and boquerones (an 

appetizer of fish, often anchovies, macerated in vinegar) in Spain 

has been implicated in infections of anisakiosis (Audicana and 

others, 2002).

Traditionally, the Japanese have consumed raw fish (sashimi) and 

raw squid (ikasashi) as a delicacy for many years (Ishikura and 

others, 1992). Gastrointestinal lesions in humans were documented in Japanese medical journals in the early 1940s (Oshima, 

1972). In a few cases small “Ascaris-like” larvae were reported, 

but it was not until 1965, after the Dutch anisakiosis cases were 

published, that the first authoritative case of human anisakiosis 

in Japan was described (Asami and others, 1965). Since then 

thousands of cases have been reported in Japan (Oshima, 1972; 

Ishikura and others, 1992; Sugimachi and others, 1985; 

Nagasawa, 1993).
Causative Agent 3

Causative Agent

Although larval anisakines were known in marine fish 

for a long time (Myers, 1976), it was not until the mid-1800s 

that nematodes causing lesions in the stomachs of cetaceans 

were grouped under the genus name Anisakis by Félix Dujardin. Initially, larval nematodes from the first human cases of 

anisakiosis in Holland were identified as Eustoma rotundatum, 

later changed to Anisakis marina, and then finally changed to 

Anisakis simplex.

The Anisakis genus, which at one time included 21 species, was reviewed in the 1960s, and 3 valid species were 

accepted based on studies of the parasite’s form and structure 

(morphology): Anisakis simplex, Anisakis typica, and Anisakis physeteris (Davey, 1971). Over the past three decades, 

the number of species has increased based on examination 

of material from intermediate and definitive hosts and the 

use of molecular and genetic methodologies (see Mattiucci 

and Nascetti, 2008, for review). Anisakis simplex sensu lato

broadly refers to a group of three sibling species, including 

Anisakis simplex sensu stricto specifically, Anisakis berlandi, 

and Anisakis pegreffi. In addition to A. typica and A. physeteris, at least five other nominal species have been described 

(table 1). It is very likely that further research will reveal new 

species in cetaceans not yet examined for parasites and expand 

the host range of known species of Anisakis (table 3).

Parasitic worms from the stomachs of seals were first 

described by Harald Krabbe in 1878 as Ascaris decipiens. 

The species was then taxonomically moved into and out of 

the genera, Porrocaeum, Terranova, Phocanema, and finally 

into Pseudoterranova. Recent research using molecular and 

genetic methodologies (see Paggi and others, 1991; Brattey 

and Davidson, 1996; Paggi and others, 2000; Zhu and others, 2002; Mattiucci and Nascetti, 2008 for reviews) has also 

determined that Pseudoterranova decipiens sensu lato consists 

of at least six sibling species: Pseudoterranova decipiens 

sensu stricto, P. krabbei, P. bulbosa, P. azarasi, P. cattani, and 

P. decipiens E (not given a name at present). Two other species were described, P. kogiae in the pygmy sperm whale and 

P. ceticola in the dwarf sperm whale (Johnston and Mawson, 

1939; Deardorff and Overstreet, 1981) (table 4). The definitive 

host range of known species of Pseudoterranova is shown in 

table 4, but future work may identify new pinniped hosts or 

new species of Pseudoterranova.

Some species of Anisakis and Pseudoterranova are found 

in the same geographic area and infect the same individual 

definitive host, indicating inbreeding, which should not occur 

if a species is biologically valid. In a few of these cases, 

hybrids of species of Anisakis and hybrids of species of Pseudoterranova are found, suggesting the taxonomic status of 

these species is still evolving (Paggi and others, 1991; Abollo 

and others, 2003; Martín-Sánchez and others, 2005; Mattiucci 

and others 2004; Mattiucci and others, 2014).

Squid

Salmon

Sashimi
4 Anisakiosis and Pseudoterranovosis

Table 3. Marine mammal hosts and geographic range of Anisakis spp.

[Species listed are those identified and described based on molecular identification except where indicated. See D`Amelio and others (2000); Mattiucci and 

Nascetti (2008); Mattiucci and others (2009); Iniguez and others (2009); Colón-Llavina and others (2009); Mattiucci and others (2014)]

Species of Anisakis Host Geographic range

Anisakis simplex sensu stricto Minke whale Northeast Atlantic Ocean.

Common dolphin East Atlantic Ocean (Iberian coastal waters).

Long-finned pilot whale Atlantic Ocean (Iberian coastal waters, South African coastal waters).

White beaked dolphin Northeast Atlantic Ocean.

Killer whale Northeast Pacific Ocean.

False killer whale Northeast Pacific Ocean.

Striped dolphin East Atlantic Ocean (Iberian coastal waters).

Beluga Northwest Atlantic Ocean.

Harbor porpoise Northeast Pacific Ocean.

Anisakis berlandi Long-finned pilot whale Southeast Atlantic Ocean (South African coastal waters), Southeast 

Pacific Ocean (New Zealand coastal waters).

Northern right whale dolphin Northeast Pacific Ocean.

False killer whale Northeast Pacific Ocean.

Southern elephant seal Sub-Antarctic waters.

Northern elephant seal Northeast Pacific Ocean.

Anisakis brevispiculata Pygmy sperm whale Atlantic Ocean (Florida, coastal waters, Iberian coastal waters, South 

African coastal waters).

Dwarf sperm whale Caribbean Sea.

Anisakis nascetti Gray's beaked whale Southwest Pacific Ocean (New Zealand).

True's beaked whale Southeast Atlantic Ocean (South African coastal waters).

Strap-toothed beaked whale Southwest Pacific Ocean (New Zealand coastal waters).

Andrew's beaked whale Southwest Pacific Ocean (New Zealand coastal waters).

Anisakis paggiae Dwarf sperm whale Atlantic Ocean (Florida coastal waters), Southeast Atlantic Ocean (South 

African coastal waters), Caribbean Sea.

Anisakis pegreffi Mediterranean monk seal1 Mediterranean Sea.

Common dolphin Atlantic Ocean (Iberian coastal waters), Western Mediterranean Sea.

Long-finned pilot whale Southeast Pacific Ocean (New Zealand coast).

Striped dolphin Mediterranean Sea.

Bottlenose dolphin Central Mediterranean Sea, South African coastal waters.

Pygmy right whale Southeast Atlantic Ocean (South African coastal waters).

Anisakis physeteris Sperm whale Central Mediterranean Sea.

Anisakis schupakovi1 Caspian seal Caspian Sea.

Anisakis typica Short-finned pilot whale Atlantic Ocean (Florida coastal waters).

Striped dolphin East Mediterranean Sea.

Bottlenose dolphin Atlantic Ocean (Florida coastal waters), Caribbean Sea.

Tucuxi Southwest Atlantic Ocean (Brazilian coastal waters).

Costero Southwest Atlantic Ocean (Brazilian coastal waters).

Pantropical spotted dolphin Northwest Atlantic Ocean (Florida coastal waters), Caribbean Sea.

Spinner dolphin Southwest Atlantic Ocean (Brazilian coastal waters).

Rough-toothed dolphin Caribbean Sea.
Causative Agent 5

Table 3. Marine mammal hosts and geographic range of Anisakis spp.

[Species listed are those identified and described based on molecular identification except where indicated. See D`Amelio and others (2000); Mattiucci and 

Nascetti (2008); Mattiucci and others (2009); Iniguez and others (2009); Colón-Llavina and others (2009); Mattiucci and others (2014)]

Species of Anisakis Host Geographic range

Anisakis ziphidarum Strap-toothed beaked whale Southeast Atlantic Ocean (South African coastal waters).

Blainville's beaked whale Southeast Atlantic Ocean (South African coastal waters).

Gervais' beaked whale Caribbean Sea.

Cuvier's beaked whale Central Mediterranean Sea, South African coastal waters, Caribbean Sea.

Gray's beaked whale Southeast Atlantic Ocean (South African coastal waters).

True's beaked whale Southeast Atlantic Ocean (South African coastal waters).

Andrew's beaked whale Southwest Pacific Ocean (New Zealand coastal waters).

1.Based on morphology only.

1.Based on morphology only.

Table 4. Marine mammal hosts and geographic range of Pseudoterranova spp.

[Species listed are those identified and described based on molecular identification except where indicated. See Johnston and Mawson (1939); Deardorff and 

Overstreet (1981); Nadler and others (2005); Mattiucci and Nascetti (2008)]]

Species of Pseudoterranova Host (common name) Geographic range

Pseudoterranova decipiens sensu stricto California sea lion Northeast Pacific Ocean.

Harbor seal Northeast Pacific Ocean.

Harbor seal Northeast Atlantic Ocean.

Harbor seal Northwest Atlantic Ocean.

Gray seal Northeast, Northwest Atlantic Ocean.

Hooded seal Northeast, Northwest Atlantic Ocean.

Northern elephant seal Northeast Pacific Ocean.

Pseudoterranova decipiens E Weddell seal Antarctic coastal waters.

Pseudoterranova azarasi Northern (Steller) sea lion Northwest Pacific Ocean, Japan Sea.

California sea lion Northwest Pacific Ocean.

Harbor seal Northeast Pacific Ocean.

Bearded seal Northwest Pacific Ocean, Japan Sea.

Pseudoterranova bulbosa Bearded seal Northeast Pacific Ocean, Northeast Atlantic Ocean, 

Northwest Atlantic Ocean, Labrador Sea.

Pseudoterranova cattani South American sea lion Southeast Pacific Ocean.

Pseudoterranova ceticola1 Dwarf sperm whale Gulf of Mexico (Mississippi).

Pseudoterranova kogiae1 Pygmy sperm whale Southwest Pacific Ocean (Queensland, Australia coastal waters), 

Southeast Atlantic Ocean (South African coastal waters).

Pseudoterranova krabbei Harbor seal Northeast Atlantic Ocean.

Gray seal Northeast Atlantic Ocean.
6 Anisakiosis and Pseudoterranovosis

Geographic Distribution

Species of Anisakis and Pseudoterranova are widely distributed in the marine environment, but most reports are from 

the Northern Hemisphere and probably reflect research interest. Most human cases of anisakiosis and pseudoterranovosis 

are from Japan, Europe, and North America; however, recent 

work in the Southern Hemisphere reports these parasites in 

various intermediate and definitive hosts as well as in humans 

(Palm, 1999; Mercado and others, 2001; Berόn-Vera and others, 2007; Torres and others, 2007). The reported low prevalence or absence of Anisakis spp. in Antarctic fish may be 

related to the distribution of definitive hosts and environmental conditions such as temperature (Measures, 1996; Klimpel 

and others, 2010). It is safe to assume that both Anisakis and 

Pseudoterranova can be found wherever suitable hosts live 

and favorable environmental conditions permit. Consequently, 

humans who consume improperly prepared infected wild fish 

or squid are at risk, especially people with cultural culinary 

practices that do not kill larvae. 

Patterns and Trends

Although cases of anisakiosis fell in Holland after legislative measures were implemented to protect consumers of fish, 

anisakiosis is still reported in Europe (Smith, 1999; Eskesen 

and others, 2001; Johansson and others, 2001; Alonso-Gómez 

and others, 2004), numbering approximately 500 human cases 

to date (Audicana and others, 2002). In Japan, more than 

2,000 cases are reported by physicians annually (Kagei and 

others, 1995). In Canada and the United States, some 50 cases 

of “anisakiasis” have been reported (McKerrow and others, 

1988), although the total may now be approaching 300 (Amin 

and others, 2000). An increasing number of cases are reported 

from South America (Mercado and others, 2001; Torres and 

others, 2007). A small number of cases of anisakiosis are 

documented from native peoples in the north (Alaska, Canada, 

Greenland), but cases are probably under-reported and misdiagnosed (Food Safety Network, 2009; Bhat and Cleland, 2010; 

Jenkins and others, 2013).

Increased reports of anisakidosis in Japan since 1972 

have been attributed to improved medical diagnostic methods such as gastroendoscopy and radiology, in addition 

to increasing populations of marine mammals, particularly 

pinnipeds in northern Japan (Oshima, 1987). In addition, 

associations between the prevalence of sealworm infections 

in commercial fish and seal populations are not fully understood (McClelland, 2002). Other abiotic factors, such as water 

temperature, are important in transmission (McClelland, 1982, 

1990, 1995; Measures, 1996; McClelland and Martell, 2001; 

McClelland, 2002). 

When “sushi bars” became popular in North America 

during the mid-1970s, cases of anisakidosis appeared, but 

pseudoterranovosis was reported more commonly than 

anisakiosis (Little and MacPhail, 1972; Little and Most, 

1973; Kates and others, 1973; Chitwood, 1975; Lichtenfels 

and Brancato, 1987; Margolis, 1977; Kliks, 1983; Desowitz, 

1986; Oshima, 1987; Amin and others, 2000). Eleven cases of 

anisakiosis and 14 cases of pseudoterranovosis were reported 

in North America between 1974–87 (Oshima, 1987). The large 

populations of pinniped definitive hosts of the parasites on 

the Atlantic and Pacific coasts of North America, as well as 

cultural dietary differences, may explain some of these cases. 

Dozens of cases of pseudoterranovosis due to consumption 

of raw fish (ceviche, also spelled cebiche) have been reported 

from Chile, Brazil, and Mexico (Torres and others, 2007). 

Many cases, however, probably go unreported or are incorrectly diagnosed. The ease of removing Pseudoterranova spp. 

from the person’s stomach by endoscopy for identification 

and the frequency by which the worm is coughed or spit out 

by the human consumer probably accounts for the increasing 

number of reports of pseudoterranovosis. Many sealworms are 

removed during commercial processing of fish or are seen by 

the consumer before consumption, thus many infections are 

likely avoided. In Atlantic Canada it has been estimated that 

annual costs to the fishing industry to remove sealworms and 

losses due to reduced value of fish products are $27–50 million Canadian dollars (see McClelland, 2002). 

Increasing populations of whales and seals, changes in 

their distribution or that of their prey, and enhanced parasite 

transmission due to favorable environmental factors, combined with the growing popularity of sushi, sashimi, and 

ceviche, are expected to raise the risk of human exposure to 

anisakids and the frequency of diagnosed anisakidosis (due to 

improved medical diagnostics).

Species Susceptibility

Humans and other nondefinitive hosts are accidentally 

infected by ingesting infected fish or invertebrates. Because 

Anisakis spp. and Pseudoterranova spp. do not mature and 

reproduce in humans or other nondefinitive hosts, these hosts 

may be considered dead-end hosts even though they are susceptible to infection. 

Human Infections

Four forms of human anisakidosis (anisakiosis and pseudoterranovosis) have been described (Smith, 1999) (table 5). 

The most commonly reported form is gastric or intestinal anisakidosis in which larvae penetrate the gastrointestinal tract, 

causing severe damage, pain, and discomfort.

The least invasive of these forms is transient luminal

anisakidosis, in which larvae do not penetrate tissues but may 

remain in the open spaces (lumen) of the gastrointestinal tract 

for some time, causing mild, temporary discomfort. These 

mild cases, while not uncommon, typically are caused by 
Box Title Here

A Case of Human Anisakiosis 7

A Case of Human Anisakiosis 

A man purchased a fresh wild salmon from a local grocery store. 

The salmon had been caught in the Pacific Ocean near the coast 

of British Columbia, Canada, gutted, and shipped whole on ice 

by air to Québec, Canada where the man lived. He consumed 

some raw salmon while cutting fillets to make marinated salmon. 

Within 24 hours, he experienced constipation, nausea, and acute 

abdominal pain, and after 2 days he went to a hospital emergency clinic. Blood samples were taken and x-rays and CT scans 

performed. Tests revealed he had a slightly above normal number 

of eosinophils, a type of white blood cell sometimes associated 

with parasitic infections, and distension and thickening of part 

of the wall of the small intestine. The man was diagnosed with 

acute abdominal syndrome, and surgery was performed. The 

surgeon removed a 15-centimeter (cm) section of thickened small 

intestine (figure). The man was discharged from the hospital 6 

days after surgery. Laboratory analysis of the resected intestine 

revealed the presence of minute abscesses (arrows) containing sections of a nematode, later identified as a third-stage 

larval Anisakis spp. This human infection is typical of intestinal 

anisakiosis. Refer to Couture and others (2003) for further details 

on this case.

Section of thickened small intestine removed during surgery. 

The arrows point to minute abscesses containing sections of 

third-stage larval Anisakis spp. (Photo by Dr. Christian Couture, 

Hôpital Laval)

Table 5. Forms of human anisakidosis.

Form

occurrence Targeted location penetration disease effects

Gastric or intestinal Most common Gastrointestinal tract Yes Mild to severe.

Outside of the Relatively rare Penetrates the wall of the gastrointes- Yes Severe.

gastrointestinal tract tinal tract and enters the body cavity

Mouth and pharynx Rare Mouth and pharynx cavity Yes Mild.

(throat)

Transient luminal Common Inner or open space of the gastrointes- No No symptoms to 

Relative Tissue Degree of 

tinal tract mild.

Pseudoterranova spp. In contrast, Anisakis spp. are 

generally more invasive and disruptive than Pseudoterranova spp. and are commonly associated with 

parasites both within and outside of the stomach 

and intestines. Most human infections are caused by 

third-stage larvae, rarely fourth-stage larvae (see

“Disease Ecology”). Symptoms and observable signs 

are described in the section “Obtaining a Diagnosis.”
8 Anisakiosis and Pseudoterranovosis

Marine Mammal Infections

The natural definitive hosts of Anisakis spp. are cetaceans. Based on the literature, at least 53 cetacean species are 

reported infected with Anisakis spp. Recently, 28 cetacean 

species infected with Anisakis spp. have had identifications 

of Anisakis to the species level based on molecular analyses 

(table 3). Immature Anisakis stages are often found in pinnipeds, where they rarely mature (Brattey and Stenson, 1993; 

Machida, 1969 cited in Kuramochi and others, 1996; Klimpel 

and others, 2008). Anisakis schupakovi in the Caspian seal 

is considered a valid species based on morphology alone 

(D’Amelio and others, 2000). Similarly, Pseudoterranova

spp. are reported as parasites of at least 28 pinniped species, 

and 11 pinniped species with Pseudoterranova spp. have had 

identifications of Pseudoterranova to the species level based 

on molecular analyses (table 4). Immature Pseudoterranova

spp. stages are occasionally found in cetaceans and sea otters, 

where they rarely mature (Measures and others, 1995; Smith, 

1999). Two species, P. kogiae in pygmy sperm whales and P. 

ceticola in dwarf sperm whales, are known based on morphology alone. 

Evidence suggests that some definitive hosts differ in 

how susceptible they are to the parasites, the number of worms 

they harbor (intensity of infection), the ability of the parasites 

to grow and reproduce in them, and their responses to Anisakis spp. or Pseudoterranova spp. For example, based on field 

data from the North Atlantic Ocean and experimental work, 

grey seals infected with P. decipiens have greater numbers of 

worms, less intense tissue reaction, greater parasite growth, 

greater parasite egg production, and higher worm survival than 

harbor seals infected with P. decipiens (McClelland, 1980a; 

Aspholm and others, 1995; McClelland, 2002). The numbers 

of P. decipiens within a host are low, and few mature worms 

are found in harp, hooded, and ringed seals (McClelland, 

2002). Some cetaceans such as minke whales and beluga can 

harbor large numbers of A. simplex of considerable size with 

high egg production, while fewer worms often are observed in 

porpoises or dolphins that harbor only larvae or small adults 

with comparatively low egg production (Brattey and Stenson, 

1995; Measures and others, 1995, Kuramochi and others, 

1996; Simard, 1997; Ugland and others, 2004; Berón-Vera 

and others, 2008). Infections of anisakids in marine mammals 

are influenced by the availability of infected intermediate or

transport hosts, host diet, host immune and physiological

status, whether the host lives in or migrates through inshore 

or offshore habitats, as well as seasonal and environmental 

factors. 

In the gastrointestinal tract of definitive hosts, anisakids 

are generally harmless (Stroud and Roff, 1979; McClelland, 

1980b; Smith, 1989; Babin and others, 1994; Abollo and 

others, 1998) even when present in high numbers, but severe 

damage has been reported in some hosts. Pathologic lesions 

caused by anisakids in marine mammals were reviewed by 

Smith (1999). Generally, anisakids provoke gastric craterlike 

ulcers or erosions where they attach to the mucous membranes 

(mucosa) in clusters of dozens to hundreds (figs. 1, 2).

In cetaceans, Anisakis spp. are found primarily in the first 

chamber of the stomach (forestomach), sometimes in the second chamber (main stomach), and rarely in the third chamber 

(pyloric stomach). Worms in the intestine are indicative of 

aberrant migrations, failed infections, or aging of the worm. 

Inflammation of the stomach (gastritis) and the intestine 

(enteritis) due to Anisakis spp. and Pseudoterranova spp. have 

been reported. In extreme cases, worms may penetrate the gut, 

enter the body cavity, and cause hemorrhage, secondary bacterial infections, inflammation of the covering of the stomach, 

intestines, and nearby organs (peritonitis), and death (Ridgway and others, 1975; Stroud and Dailey, 1978; Martineau and 

others, 1988; Baker and Martin, 1992; De Guise and others, 

1995; Lair and others, 2014).

In unusual hosts, such as the sea otter, P. decipiens

(primarily larvae) is associated with stomach ulcers, holes 

or perforations in the intestines, and death due to peritonitis 

(Rausch, 1953; Fay and others, 1978 in Smith, 1999). Lesions 

in the pyloric region of the simple stomach may cause narrowing, physically preventing the movement of food or leading to 

an abnormal accumulation of blood in the veins (Smith, 1999). 

Ulceration and gut perforation by parasites such as A. simplex

can provide portals of entry for viruses or bacteria, some 

of which may lead to abscesses, systemic infections of the 

entire body, or possibly other chronic diseases (Martineau and 

others, 1988; Baker and Martin, 1992; De Guise and others, 

1994, 1995; Harper and others, 2002a, 2002b; Lair and others, 

2014).

Fish and Invertebrates

Larval Anisakis spp. have been reported from a variety 

of naturally infected invertebrate intermediate hosts, mostly 

crustaceans such as small shrimplike crustaceans (euphausiids and amphipods), crayfish, crabs, lobsters, and shrimp 

(decapods), and squid and cuttlefish (cephalopods). Larval 

Pseudoterranova spp. have been reported in amphipods, 

decapods, and small, shrimplike crustaceans (mysids) (Uspenskaya, 1963; Oshima and others, 1968; Smith, 1971; Nagasawa and Moravec, 1995; Hays and others, 1998; McClelland, 

2002). P. decipiens is known from over 75 fish species belonging to 29 families, 10 orders, and 3 classes of fish (McClelland, 2002). Anisakis spp. are reported from a wide range of 

marine, estuarine, and occasionally freshwater fish (Oshima, 

1972; Smith and Wootten, 1978) and occur in approximately 

200 species of fish and 25 species of cephalopods worldwide 

(Klimpel and others, 2004). However, these figures may be 

inaccurate due to confusion in larval identification (Oshima, 

1972; Smith and Wootten, 1978). For example, one report 

states that one Anisakis “larval type 1” is known from 123 

different fish species in Japanese waters (Oshima, 1972), yet 

another report states that more than 166 species of fish are 

infected with anisakids in Japanese waters (Ishimura and 

others, 1992).
Species Susceptibility 9

Figure 1. Adult Anisakis simplex in the stomach of a pilot whale. 

(Photo by Dr. Lena Measures, Fisheries and Oceans Canada)

Figure 2. Adult Pseudoterranova decipiens in the stomach of a grey seal. (Photo by Dr. Lena Measures, 

Fisheries and Oceans Canada)
10 Anisakiosis and Pseudoterranovosis

Freshwater fish are likely to become infected if they are 

fed the infected internal organs of marine fish in aquacultural 

operations or if wild fish such as salmon spend part of their 

life cycle in salt or estuarine waters where they may acquire 

infections. Seven cephalopod species can act as transport hosts 

of A. simplex and A. physeteris in Japanese waters (Nagasawa, 

1993), and seven cephalopod species have been infected with 

A. simplex in Spanish waters (Abollo and others, 1998). In 

cephalopods, Anisakis spp. larvae are found in the internal 

organs and muscles, and in a few cases, destroy the testes or 

ovaries of mature squid (Abollo and others, 1998). Although 

Anisakis spp. and Pseudoterranova spp. are reported from 

a wide range of fish hosts, it is believed that primarily fish 

that live in the water column (pelagic fish) are ecologically 

important for transmitting Anisakis spp. to cetaceans, and fish 

that live near or on the bottom (benthic fish) are ecologically 

important for transmitting Pseudoterranova spp. to pinnipeds.

To understand the role of invertebrates and fish in the life 

cycles of these two anisakids, various experimental infections 

have been conducted since the early 1950s. For example, 12 

different species of fish (marine, anadromous, and freshwater) including Atlantic salmon, brook trout, rainbow smelt, and 

Atlantic cod were susceptible to experimental infections by P. 

decipiens using experimentally infected amphipod intermediate hosts (McClelland, 1995). In some fish such as rainbow 

smelt, tissue responses to infections were absent. In other 

fish, larvae were surrounded or encapsulated by cells of the 

immune system and eventually killed. Anisakis spp. larvae are 

rapidly encapsulated in fish, and some larvae die and become 

hardened by deposited calcium salts (Smith, 1974). Although 

Anisakis spp. appear to cause few pathologies in fish—unless 

the liver is heavily infected or there are large numbers of 

worms—sealworms can kill small fish, particularly when vital 

organs are damaged by migrating larvae (McClelland, 1995). 

Experimental evidence indicates that P. decipiens infection of 

some fish species affects their ability to swim and thus escape 

predators (Sprengel and Lüchtenberg, 1991).

An unusual condition called red-vent syndrome (RVS; 

fig. 3), associated with heavy infections by larval Anisakis

spp. in the skin and muscle tissues of the vent area of wild 

Atlantic salmon, has been reported in the North Atlantic (Beck 

and others, 2008) including recently in Canada (Larratt and 

others, 2013). Lesions appear as small red spots (small hemorrhages) with reddening around the vent in light infections 

to pronounced swelling, hemorrhage, erosion of the skin’s 

outer layer, loss of scales, and skin inflammation (dermatitis) in severe infections. It is currently unknown why this 

occurs, but RVS may be an external sign of heavily parasitized fish (Murphy and others, 2010). However, the intensity 

of larval Anisakis infections in wild Atlantic salmon was only 

weakly associated with RVS, suggesting other factors may 

be involved in this syndrome such as immune status, stress, 

water temperature and timing 

of infections (Larrat and others, 

2013). Increased levels of infections have been observed in adult 

salmon returning to fresh water 

to spawn (Murphy and others, 

2010). Apparently, after Atlantic 

salmon spawn, red vent lesions 

subside somewhat. There is no 

evidence that RVS causes mortality or prevents salmon from 

spawning but further research is 

warranted. 

Figure 3. Red-vent syndrome 

in Atlantic salmon. (Photo by 

Ministère des resources naturelles 

et de la faune [Station piscicole de 

Tadoussac]).
Obtaining a Diagnosis 11

Birds

There are a few records of Anisakis spp. larvae in fisheating birds, particularly in Japanese waters. Larval Anisakis

spp. have been reported from the herring gull, black-tailed 

gull, tufted puffin, Swinhoe’s storm petrel, Japanese cormorant, and northern fulmar (Riley, 1972; Smith and Wootten, 

1978). A single immature adult female Anisakis spp. was 

reported from the esophagus of a northern fulmar in northern 

Europe (Riley, 1972). The occurrence of these parasites in 

fish-eating birds is regarded as accidental.

Other Animals 

Experimentally or inadvertently, frogs, turtles, cats, dogs, 

rabbits, rats, guinea pigs, domestic pigs, and monkeys were 

susceptible to infection with Anisakis spp. or Pseudoterranova

spp., but results varied (Smith and Wootten, 1978; Smith, 

1999). In some cases, worms migrated through various tissues 

and caused pathologies; in other cases, worms died and were 

encapsulated by strong tissue responses. Natural infections of 

a brown bear (Smith and Wootten, 1978; Smith, 1999) and a 

dog (Kitayama, 1962 in Oshima, 1987) have been reported.

Obtaining a Diagnosis

Many human cases of anisakiosis and pseudoterranovosis 

likely go unreported, and probably many worms die or are 

passed in the feces without causing any symptoms or clinical 

signs. Diagnosis is difficult, but clinical infections are usually suspected based on symptoms, clinical presentation, and 

history of recent consumption of improperly prepared seafood. Symptoms in acute cases of gastric anisakidosis include 

intense upper abdominal pain, nausea, and vomiting within 5 

hours of consuming infected fish or squid.

Intestinal anisakidosis has less specific symptoms, but 

usually the victim experiences low abdominal pain, particularly in the lower right quadrant; nausea; vomiting; fever; 

diarrhea or constipation; and inapparent blood in the stool, 

usually within 1–7 days after consuming infected fish or squid. 

Infections, either gastric or intestinal, may become chronic, 

and intestinal blockage and tumors (swellings) have been 

reported (Smith and Wootten, 1978; Smith, 1999). Transient 

infections with Pseudoterranova spp. are often diagnosed 

based on the reported symptom of a “tingling” sensation in the 

throat or mouth or mild abdominal discomfort, and the worms 

are vomited or coughed up by the patient days to weeks after 

ingestion (Smith, 1999).

In clinical cases of gastric infection, Pseudoterranova

spp. may be diagnosed using gastroendoscopy, whereby a 

fiberscopic tube equipped with a small camera and forceps is 

passed through the person’s mouth into the stomach to observe 

the surface of the mucous membranes (Oshima, 1987). Worms 

can then be removed by using fine biopsy forceps. The same 

method can be used for gastric infections of Anisakis spp. 

(Bhat and Cleland, 2010), but not for most intestinal infections 

due to the physical limit of endoscopy. Fifty-six larval Anisakis spp. were removed by gastroendoscopy from the stomach 

of a Japanese patient who had prepared her own sashimi 

(Kagei and Isogaki, 1992). Other medical imaging such as

radiology, computed tomography (CT) scans, and abdominal ultrasound may help in diagnoses (Oshima, 1987; Couture 

and others, 2003). 

Blood and blood serum tests such as enzyme-linked 

immunosorbent assay (ELISA) have been used to detect 

specific immunoglobulin E (IgE) antibodies to A. simplex (Lorenzo and others, 2000; Valiñas and others, 2001) 

in blood samples. Histological analysis of biopsied lesions 

caused by Anisakis spp. or Pseudoterranova spp. attaching to 

the gastrointestinal wall usually contain eosinophils, a type 

of white blood cell indicative of parasitic infection. Severe 

allergic reactions such as dermatitis, subcutaneous swelling, 

rashes, throat swelling, and low blood pressure, particularly 

in response to A. simplex, have been reported in acute cases, 

especially in Spain where more than 150 cases have been 

reported since 1995 (Audicana and others, 2002). In these 

cases, specific IgE and skin prick tests have been used in 

diagnosing allergic reactions to A. simplex, but acute parasitic 

infections and allergies are still under investigation (AlonsoGόmez and others, 2004; Reddy and Fried, 2008).

Where clinical infections are rarely encountered by the 

medical profession, such as in North America and Europe, 

infections may be misdiagnosed as gastrointestinal cancer, 

ulcers, obstruction or inflammation such as Crohn’s disease, 

appendicitis, and acute abdominal syndrome. Consequently, 

abdominal surgery is often performed. It is during surgery 

that worms, often alive, may be found (Clavel and others, 

1993), and usually affected tissues are removed. Histological, 

morphological, and molecular analyses can be used to identify 

the cause of tissue inflammation as well as identify the genus, 

and sometimes the species, of recovered worms (Maejima and 

others, 1992; Eskesen and others, 2001; Couture and others, 

2003; Mattiucci and Nascetti, 2008). Identification of larval 

Anisakis spp. and Pseudoterranova spp. is based principally 

on morphology of the digestive system of the worm studied 

using light microscopy (fig. 4).
12 Anisakiosis and Pseudoterranovosis

Figure 4. Key morphological features of anisakid larvae removed from fish: A, Anisakis; B, Contracaecum/Phocascaris; 

C, Pseudoterranova. Note proventriculus (p), ventriculus (v), intestine (i), intestinal caecum (ic), ventricular appendage (va), boring tooth (bt), 

excretory pore (ep), nerve ring (nr), excretory duct (ed), excretory canal (ec).

A. Head-Anisakis B. Head-Contracaecum/Phocascaris C. Head-Pseudoterranova

ep

nr

1 mm

1 mm

ed

ic

p

v

i

ec

va

bt

ep

nr

ed

p

v

i

ec

bt

ep

nr

ed

p

v

i

ec

ic

bt

1 mm

Figure 5. Key morphologic features of anisakid larvae ...
Disease Ecology 13

Disease Ecology

The life cycle of Anisakis spp. is complex and involves 

pelagic invertebrates such as small shrimplike crustaceans or 

euphausiids (commonly called krill) or squid and pelagic fish 

such as capelin, herring, salmon, or redfish as intermediate 

or transport hosts, with cetaceans as definitive hosts. The life 

cycle of Pseudoterranova spp. is similar but involves benthic 

invertebrates such as amphipods (commonly called scud), and 

benthic fish such as flatfish and cod as intermediate hosts, with 

pinnipeds as definitive hosts.

Anisakis spp. reproduce in the stomach of cetaceans such 

as minke whales and beluga and harbor porpoises (fig. 5). 

Female worms produce eggs, which enter the sea with the 

feces of the cetacean. A period of development is required 

in the sea before larval stages hatch from eggs. These larvae 

are then eaten by a necessary, or “obligate,” first intermediate 

host such as krill, where larvae develop and grow considerably, reaching 7– 39 millimeters (mm) in length. Infected krill 

are subsequently eaten by a transport host such as squid or 

small pelagic fish (capelin, mackerel, herring). These infected 

fish or squid are in turn eaten by large pelagic predatory fish 

such as redfish, salmon, or silver hake, which can accumulate 

large numbers of infective larvae. Finally, the life cycle is 

completed when the definitive cetacean host eats fish (small 

or large fish) or krill containing infective larvae. Larval stages 

Figure 5. The life cycle of Anisakis simplex. (Adapted from Anderson, 2000)

Larvae mature to adults

in cetacean stomach.

Eggs enter sea with 

feces of cetacean.

Larvae develop and

hatch from eggs.

Free-swimming 

larvae eaten by 

krill.

Infected krill eaten 

by squid and small fish.

Larvae are transferred 

fish-to-fish through predation.

Humans are accidentally infected by 

eating raw or undercooked seafood.

Cetaceans ingest infective larvae 

by eating krill, squid or fish. 
14 Anisakiosis and Pseudoterranovosis

in fish are small (9–39 mm long) and are usually found in a 

flat coil on the visceral organs (fig. 6). They may also occur in 

the flesh of fish where they are difficult to see, because they 

are transparent or white in color. Larvae may migrate from the 

visceral organs into the muscle after death, especially if the 

fish is allowed to warm to room temperature. 

Pseudoterranova spp. reproduces in the stomach of 

pinnipeds such as grey seals (fig. 7). Female worms produce 

eggs that enter the sea with the feces of the seal. A period of 

development is required in the sea before larval stages hatch 

from eggs. These larvae attach to sand or rocks by their tails 

and are eaten by a first host such as copepods or immature or 

mature invertebrates such as mysids, amphipods, isopods, and 

polychaetes. Infected invertebrates are subsequently eaten 

by a second host such as small benthic fish (sculpins, smelt, 

juvenile cod, and flatfish). These small infected fish are in 

turn eaten by large benthic predatory fish such as large cod, 

monkfish, and eelpout, which can accumulate large numbers 

of infective larvae. Finally the life cycle is completed when 

the definitive seal host eats small or large fish containing 

infective larvae. The biology of Pseudoterranova spp. is not 

completely understood, particularly the role of invertebrates 

and small fish (McClelland, 2002; Smith and Hemmingsen, 

2003). Larval stages in fish grow considerably, from more than 

2– 60 mm long, depending on the size of fish and its age when 

it is infected. Larvae may also reside in the gut wall, liver, or 

body cavity of fish, but most are found in the muscles (fig. 8). 

As larvae are yellow to red in color and relatively large, they 

are often seen in the fillets, much to the alarm of consumers. 

Larvae may or may not be encapsulated and may lie coiled or 

fully extended in tissues, but this may be a function of host 

species, temperature, or age of infection (Ramakrishna and 

Burt, 1991; McClelland, 1995). Pseudoterranova spp. larvae 

may also migrate after the death of the fish. Similarly, another 

related anisakid, Hysterothylacium aduncum, which matures in 

the intestine of fish such as cod, can migrate after the host dies 

by exiting from its anus, mouth, or gills and is often misidentified as Pseudoterranova spp.

Figure 6. Larval Anisakis sp. (arrows) in the abdominal cavity of an Atlantic herring. (Photo by Dr. Lena Measures, 

Fisheries and Oceans Canada)
Disease Ecology 15

Figure 7. The life cycle of Pseudoterranova decipiens. (Adapted from McClelland, 2002, and Anderson, 2000)

Figure 8. Larval Pseudoterranova sp. in 

muscle of an American plaice. (Photo by 

Dr. Lena Measures, Fisheries and Oceans 

Canada)

Larvae mature to adults

in seal stomach.

Seals ingest infective 

larvae by eating fish.

Eggs enter sea with feces of seal.

Larvae develop and

hatch from eggs.

Larvae attach to sand 

or rocks.

Larvae eaten by 

invertebrate host.

Infected invertebrate 

eaten by small fish.

Larvae are transferred 

fish-to-fish through predation.

Figure 8. Life cycle of Pseudoterranova decipiens.

Humans are accidentally infected by 

eating raw or undercooked seafood.
16 Anisakiosis and Pseudoterranovosis

Migration of Adult Hysterothylacium aduncum in Atlantic Cod 

After Host Death

Post-mortem migration of nematodes is a well recognized 

phenomenon observed by parasitologists, fishermen, hunters, 

wildlife biologists, and others. It is particularly common in coldblooded animals such as fish when the temperature of a recently 

caught fish increases above that of its original habitat in water. 

The rise in temperature stimulates some parasites in fish tissues 

to migrate within the body of the fish or even to exit the body 

completely. In the figure below, Hysterothylacium aduncum, a 

parasitic nematode found as adults in the digestive tract of Atlantic cod, the definitive host, leave the fish via the vent, mouth, or 

gills. Many marine fishermen are familiar with sealworm, Pseudoterranova decipiens, which uses cod as an intermediate host 

in which larval stages occur in the muscles. Although sealworm 

larvae migrate within the tissues of cod, even when cod are still 

alive, many fishermen mistake H. aduncum for sealworm when 

they observe a fish as illustrated in the figure below. Post-mortem 

migration by H. aduncum can be eliminated by gutting fresh cod 

immediately after capture.

Hysterothylacium aduncum, a parasitic nematode of Atlantic cod, may be mistaken for P. decipiens as they exit the body of a dead fish 

through the vent, mouth, or gills. (Photo by Dr. Lena Measures, Fisheries and Oceans Canada)
Commercial Candling of Fish Fillets to Visualize and Extract Pseudoterranova decipiens 17

Commercial Candling of Fish Fillets to Visualize and 

Extract Pseudoterranova decipiens

Candling of fish in fish-processing plants is commonly carried out 

in areas of the world where fish are frequently infected with parasitic nematodes such as sealworm, Pseudoterranova decipiens, 

because worms look unsavory to fish consumers. Candling allows 

the workers to see the worms and remove them from fish fillets. 

Fillets are sliced to a predetermined thickness and placed on a 

light table. Light passing through the fillet from below enables 

workers to see large, yellow to red larval sealworms and then 

remove them using forceps. As the efficacy of detection and 

removal depends on various factors such as the thickness of the 

fish fillet, the number of worms present, their size, the intensity of 

light used, and observer experience, candling is not 100 percent 

efficient. Candling involves considerable cost to fish processers 

and diminishes the quality and value of fillets.

Candling of fish fillets to visualize and remove larval worms. (Photo by Dr. Lena Measures, Fisheries and Oceans Canada)
18 Anisakiosis and Pseudoterranovosis

Points to Ponder

Most professional chefs in “sushi-bars” or high quality 

restaurants that serve sashimi, ikasashi, or ceviche are trained 

in the proper handling of raw fish or squid for safe consumption by their customers. It is more commonly the home chef 

preparing his own fish dish that has an intimate experience 

with parasitic worms (Kates and others, 1973; Oshima, 1987; 

Kagei and Isogaki, 1992; Maejima and others, 1992; Couture 

and others, 2003). The risk of anisakid infection from consuming farmed fish (those raised in sea pens, cages, or sea nets in 

aquacultural operations where fish are fed a commercial food, 

usually dry or expanded pellets) is considered low (Deardorff 

and Kent, 1989; Inoue and others, 2000).

Disease Prevention and Control

Infective larval Anisakis spp. and Pseudoterranova spp. 

are remarkably resistant to various environmental conditions. In fish flesh they can survive cold smoking, insufficient 

marination or salting, and freezing temperatures (Hauck, 

1977; Bier, 1976; Deardorff and Throm, 1988; Gardiner, 1990; 

Wharton and Aalders, 2002). Several methods of fish preparation can be used to reduce the chance of acquiring infections 

(table 6). Rapid evisceration of freshly caught fish is important should there be larvae in the viscera or body cavity (Smith 

and Wootten, 1975). Larval Anisakis spp. in fish flesh is problematic for consumers, because its small size and transparent 

or white color makes it difficult to detect. On the other hand, 

larval Pseudoterranova spp., which is commonly commonly 

found in fish flesh, is often seen by the consumer, depending 

on the thickness of fish fillets. Slicing fish fillets and using a 

strong light source to visualize larvae and then remove them 

manually (a process called candling) is still used commercially 

in some fish processing plants, but candling is costly, diminishes the value of the product, and is inefficient as its efficacy 

depends on the thickness of the fillet being examined (Power, 

1958, 1961; Valdimarsson and others, 1985; Hafsteinsson and 

Rizvi, 1987). An effective research tool to detect and obtain 

live larval anisakids from infected invertebrates, fish, or other 

tissue is artificial digestion using a Baermann apparatus.

Research has shown that commercial blast-freezing of 

infected fish to at least –35 degrees Celsius (°C) for 15 hours 

will kill larvae (Deardorff and Throm, 1988). However, larval 

survival in fish flesh is dependent on the mass of the fish 

being frozen. For example, 20-kilogram (kg) boxes of monkfish required more than 28 hours to reach the commercial 

blast freezer temperature of –35 °C to –38 °C (Wharton and 

Aalders, 2002). Furthermore, it is important that all parts of 

the fish freeze solid at these temperatures and for the required 

time to ensure killing any larvae that may be present. According to the U.S. Food and Drug Administration, raw, rawmarinated, partially cooked, or marinated partially cooked fish 

must be frozen at –35 °C or below until solid and stored at this 

temperature or below for 15 hours to destroy larvae. Similarly, 

freezing these products at –20 °C or below for a minimum of 

168 hours (7 days) will also destroy larvae (U.S. Food and 

Drug Administration, 2009). Cooking fish thoroughly so that 

the internal temperature of the fish attains 60–63 °C for a few 

minutes will kill larvae (Oshima, 1972; U.S. Food and Drug 

Administration, 2009). 

Table 6. Methods to reduce risk of infection by anisakid 

larvae for consumers of fish.

[°C, degrees Celsius]

• Rapid evisceration of fish to prevent postmortem migration of 

larvae into muscles.

• Candling and manual removal of larvae.

• Solid freezing of fish. The time required to freeze fish solid 

depends on the mass of the fish. Recommended to freeze fish at 

–20 °C for a minimum of 7 days.

• Thorough cooking of fish to an internal temperature of 

60–63 °C for several minutes.
The Baermann Technique 19

The Baermann Technique

The Baermann technique is a method commonly employed in 

laboratories to concentrate and collect live organisms from a 

large sample of soil, water, or tissue. It was developed by G. 

Baermann in 1917 to recover hookworm larvae from soil samples 

(Baermann, 1917). It consists of a funnel with a plastic tube at the 

bottom that is closed with a stopcock, which can be opened to 

remove small samples or liquids containing organisms that have 

accumulated in the plastic tube. A sample of soil or tissue may be 

placed on a sieve or filter suspended in the top part of the funnel. 

A liquid medium such as water may or may not be used. There are 

various modifications of the Baermann apparatus. A light may be 

suspended above the funnel to encourage light-averse organisms 

to migrate downward, a technique frequently used by entomologists to collect some insects or soil organisms. In parasitology, 

a modified version may involve preparing a pepsin solution to 

simulate the digestive juices in the stomach of a mammal or bird. 

This solution is poured into the funnel with the stopcock closed, 

and infected tissue such as that from a fish or invertebrate is 

placed on a sieve or filter within the fluid at the top of the funnel. 

The apparatus may be kept at room temperature, 37 degrees 

Celsius (°C) or 42 °C depending on the sample. Enzymes in the 

pepsin solution slowly digest the tissue, such as fish muscle 

containing larval Anisakis simplex, and the enzymes and warm 

solution stimulate the larvae to migrate out of the tissues and fall 

downward into the plastic tube where, after a few hours incubation, they may be collected. Samples of krill can also be digested 

in this manner to collect larval Anisakis spp. (Hays and others, 

1998).

Baermann apparatus. (Photo by Dr. Lena Measures, Fisheries and Oceans Canada)
20 Anisakiosis and Pseudoterranovosis

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26 Anisakiosis and Pseudoterranovosis

Glossary

A

aberrant Departing from the usual course; 

unusual.

abiotic Nonliving.

abscess A localized collection of pus 

(thick, yellowish fluid made of white blood 

cells, dead tissue cells, and cellular debris) 

surrounded by inflamed tissue.

amphipod A small, shrimplike crustacean 

within the taxonomic order Amphipoda, having a laterally compressed body and living 

mainly in aquatic habitats. See crustacean, 

scud.

anadromous Migrating up rivers from the 

ocean to breed in fresh water.

anisakid A roundworm within the family 

Anisakidae, including members of the genera 

Anisakis and Pseudoterranova. See genera, 

taxonomic.

antibody A protein formed in the body 

of a vertebrate that is used by the immune 

system to identify and neutralize the effects 

of foreign invading proteins (antigens), such 

as those in bacteria and viruses.

ascarid Any parasitic nematode worm of 

the family Ascaridae. See parasite, nematode.

B

benthic Pertaining to the bottom of the 

ocean or lake or the organisms that live there.

C

cephalopod A mollusk within the taxonomic class Cephalopoda having a large head 

and tentacles, for example, octopus, squid, 

cuttlefish, and nautilus. See taxonomic.

cetacean A marine mammal within the 

taxonomic order Cetacea having no hind 

limbs, flippers as forelimbs, and a blowhole 

for breathing, for example, whales, dolphins, 

and porpoises. See taxonomic.

copepod A tiny, aquatic crustacean within 

the taxonomic subclass Copepoda, having an 

elongated body and a forked tail. See crustacean, taxonomic.

crustacean An invertebrate animal within 

the taxonomic class Crustacea, having a 

segmented body, chitinous exoskeleton, and 

jointed limbs, for example, lobsters, crabs, 

shrimp, barnacles, and pill bugs. See invertebrate, taxonomic.

computed tomography (CT) scan An 

imaging method used to construct a crosssectional image of body structures in a body 

plane produced by x-ray absorption of the 

structures in the image plane.

D

dead-end host A host in which a parasite 

cannot complete its development and transmission in order to continue its life cycle. 

See host, definitive host, intermediate host, 

parasite.

decapod A crustacean within the taxonomic 

order Decapoda, having 10 legs, for example, 

crab, lobster, and shrimp. See crustacean, 

taxonomic.

definitive host An organism in which sexually mature stages of a parasite occur. Also 

called the final host. See host, parasite.

dermatitis Inflammation of the skin.

E

encapsulated Enclosed within a protective 

layer of cells.

endoscopy Visual examination of the inside 

of organs and cavities of the body using an 

illuminated, flexible optical tube.

entomologist A scientist who studies 

insects.

enzyme A protein produced by a living cell 

that functions to increase or decrease the rate 

of metabolic processes.

eosinophils The white blood cells involved 

in combating parasitic infections and controlling allergic reactions. See parasitic.

estuarine Pertaining to an estuary, an area 

where a river nears the ocean and fresh water 

mixes with tidal salt water.

euphausiid A small shrimplike crustacean 

within the taxonomic order Euphausiacea. 

See crustacean, taxonomic, krill.

eviscerate To remove the internal organs; 

disembowel.
Glossary 27

F

fish Refers to finfish, which are coldblooded strictly aquatic vertebrates with a 

well differentiated skull and a bony skeleton, 

rather than shellfish (see invertebrates) and 

jawless fishes. G

gastric Pertaining to the stomach.

gastroendoscopy Endoscopy of the stom-

ach. See endoscopy.

gastrointestinal tract The tubular organs 

that form a digestive pathway from the mouth 

to the anus, including the esophagus, stom-

ach, and intestines.

genera Plural of genus. See genus.

genus A taxonomic group between family 

and species containing organisms that share 

common characteristics. See taxonomic, 

genera. H

histologic Pertaining to histology, the study 

of the microscopic structure of tissues.

host An organism that harbors or nourishes 

microbes, viruses, and parasites. See parasite. I

Immunoglobulin E (IgE) A type of antibody 

that is important in allergic reactions. See

antibody.

immune status Refers to the host’s ability 

to fight infections. See host.

intensity Number of parasites in an infected 

host. See parasite, host.

intermediate host An organism in or on 

which the larval stage of a parasite develops 

but does not sexually reproduce. See host, 

parasite, larva/larval.

Inuit Native people of northern North 

America, inhabiting areas from Greenland 

and eastern Canada to Alaska.

invertebrates Animals lacking a spinal 

column, for example, insects and crustaceans; 

vertebrates, in contrast, possess a spinal col-

umn. See crustacean.

isopod A crustacean within the taxonomic 

order Isopoda, characterized by a flattened 

body and gills located on abdominal append-

ages; most are aquatic bottom-dwellers in 

fresh or salt water. See crustacean, taxo-

nomic. JK

krill A small shrimplike crustacean within 

the taxonomic order Euphausiacea. See crus-

tacean, taxonomic, euphausiid. L

larva/larval The immature, early form of 

an organism that at birth or hatching is not 

like its parent and has to undergo a series of 

form and size changes before assuming adult 

features.

lesion An abnormal change in tissue or an 

organ due to disease or injury.

light microscopy A technique using visible 

light and single or multiple lenses (micro-

scope) to obtain a magnified view of a 

sample.

luminal Pertaining to the lumen, the inner 

open space of a tubular organ. M

morphology The study of the form and 

structure of an organism or its parts.

mucosa A mucous membrane or moist 

tissue layer lining hollow organs such as the 

stomach and cavities of the body.

mysid A small, shrimplike crustacean 

within the taxonomic orders Mysida or 

Lophogastrida found in salt and fresh water. 

See crustacean, taxonomic. N

nematode An unsegmented, cylindrical 

parasitic worm within the taxonomic class 

Nematoda; roundworm. See parasitic, taxo-

nomic. OP

parasite An organism that lives in or on 

another organism of a different species from 

which it derives nutrients and shelter.

parasitic Pertaining to parasites. See para

-

site.
28 Anisakiosis and Pseudoterranovosis

parasitologist A scientist who studies parasites. See parasite.

pelagic Pertaining to the upper waters of the 

open ocean or the organisms that live there. 

pepsin An enzyme produced in the stomach 

that breaks down proteins.

perforation An abnormal hole in an organ or 

the act of creating such a hole.

peritonitis Inflammation of the peritoneum, 

the membrane lining the abdominal cavity and 

internal organs.

physiological status Refers to the host`s 

physiology, the normal functioning of an 

organism.

pinniped A marine mammal within the 

taxonomic suborder Pinnipedia having finlike 

flippers as feet, such as walrus, sea lions, fur 

seals, and earless seals. See taxonomic.

polychaete A segmented worm, generally 

marine, within the taxonomic class Polychaeta. See taxonomic.

prevalence The total number of cases of a 

disease divided by the number of hosts examined in a population at a given time. See host.

Q

R

radiology The science of using electromagnetic radiation or sound waves to make 

images of structures within the body, including x-rays, computed tomography (CT) scans, 

magnetic resonance imaging (MRI), and ultrasound. See computed tomography scan.

roundworm See nematode.

S

scud A tiny crustacean resembling a shrimp. 

See crustacean, amphipod.

sensu lato In a broad or wide sense.

sensu stricto In a strict or narrow sense.

spawn To deposit eggs or produce offspring.

species A population or populations of 

closely related and similar organisims that 

freely interbreed with one another in natural 

conditions but not with members of other 

species.

Taxonomic hierarchy, 

from general to specific

Kingdom

Phylum

Class

Order

Family

Subfamily

Genus

Species

transport host An organism in which a 

parasite survives without further development. See parasite, host.

U

V

vent The common opening of the intestinal, genital, and urinary tracts in fish, 

reptiles, and birds. See fish.

viscera The internal organs, particularly 

of the thoracic and abdominal cavities, 

such as the heart, lungs, liver, kidneys, and 

intestines.

visceral Pertaining to viscera. See viscera.

W

X

Y

Z

zoonotic Transmissible between animals 

and humans, and vice versa.

susceptibility/susceptible Pertaining to 

the ability of an animal to become infected 

by a disease-causing organism or to 

develop disease after becoming infected.

T

taxonomic Pertaining to taxonomy. See

taxonomy.

taxonomy The systematic principles 

and procedures of grouping and arranging organisms into a hierarchical order 

(below).
Appendixes

Appendix 1. Common and Scientific Names for Species Cited

Appendix 2. Important Literature About Anisakis spp. and Pseudoterranova spp.
30 Anisakidosis and Pseudoterranovosis

Appendix 1. Common and Scientific Names for Species Cited

Common name Scientific name

Birds

Black-tailed gull Larus crassirostris

Herring gull Larus argentatus

Northern fulmar Fulmarus glacialis

Swinhoe’s storm petrel Oceanodroma monorhis

Tufted puffin Fratercula cirrhata=Lunda cirrhata

Japanese cormorant Phalacrocorax capillatus

Crustaceans

Crabs Order Decapoda

Crayfish Order Decapoda

Krill Order Euphausiacea

Amphipods Order Amphipoda

Lobsters Order Decapoda

Shrimp Order Decapoda

Fish

American plaice Hippoglossoides platessoides

Anchovy Family Engraulidae

Atlantic cod Gadus morhua

Atlantic salmon Salmo salar

Brook trout Salvelinus fontinalis

Capelin Mallotus villosus

Eelpout Family Zoarcidae

Flatfish Order Pleuronectiformes

Herring Family Clupeidae

Mackerel Scomber scombrus

Monkfish Family Lophiidae

Rainbow smelt Osmerus mordax

Redfish Sebastes scombrus

Salmon Family Salmonidae

Sculpin Family Cottoidei

Silver hake Merluccius bilinearis

Smelt Family Osmeridae
Appendix 1. Common and Scientific Names for Species Cited 31

Common name Scientific name

Mammals

Andrew's beaked whale Mesoplodon bowdoini

Bearded seal Erignathus barbatus

Beluga Delphinapterus leucas

Blainville's beaked whale Mesoplodon densirostris

Bottlenose dolphin Tursiops truncata

Brown bear Ursus arctos

California sea lion Zalophus californianus

Caspian seal Phoca caspica

Common dolphin Delphinus delphis

Costero Sotalia guianensis

Cuvier's beaked whale Ziphius cavirostris

Dog (domestic) Canis lupus familiaris

Dwarf sperm whale Kogia sima

False killer whale Pseudorca crassidens

Gervais' beaked whale Mesoplodon europaeus

Gray's beaked whale Mesoplodon grayi

Gray seal Halichoerus grypus

Harbor porpoise Phocoena phocoena

Harbor seal Phoca vitulina

Harp seal Pagophilus groenlandicus

Hooded seal Cystophora cristata

Killer whale Orcinus orca

Long-finned pilot whale Globicephala melas

Mediterranean monk seal Monachus monachus

Minke whale Balaenoptera acutorostrata

Northern elephant seal Mirounga angustirostris 

Northern right whale dolphin Lissodelphis borealis

Northern (Steller) sea lion Eumetopias jubatus

Pantropical spotted dolphin Stenella attenuata
32 Anisakidosis and Pseudoterranovosis

Common name Scientific name

Mammals—Continued

Pygmy right whale Caperea marginata

Pygmy sperm whale Kogia breviceps

Ringed seal Pusa hispida

Rough-toothed dolphin Stenella longirostris

Sea otter Enhydra lutris

Short-finned pilot whale Globicephala macrorhynchus

South American sea lion Otaria flavescens

Sperm whale Physeter macrocephalus

Spinner dolphin Stenella longirostris

Southern elephant seal Mirounga leonina 

Strap-toothed beaked whale Mesoplodon layardii

Striped dolphin Stenella coeruleoalba

True's beaked whale Mesoplodon mirus

Tucuxi Sotalia fluviatilis

Weddell seal Leptonychotes weddellii

White beaked dolphin Lagenorhynchus albirostris

Mollusks

Squid Class Cephalapoda, Order Teuthida

Cuttlefish Class Cephalapoda, Order Sepiida
Appendix 2. Important Literature About Anisakis spp. and Pseudoterranova spp. 33

Appendix 2. Important Literature About Anisakis spp. and Pseudoterranova spp.

Anisakiosis and Pseudoterranovosis

Berland, B., and Fagerholm, H.-P., 1994, Thirty years of 

anisakid research, and future trends: Bulletin of the 

Scandinavian Society of Parasitology, v. 4, p. 40–46. 

Cheng, T.C., 1976, The natural history of anisakiasis in 

animals: Journal of Milk and Food Technology, v. 39, 

p. 32–46.

Ishikura, H., Kikuchi, K., Nagasawa, K., Ooiwa, T., Takamiya, H., Sato, N., and Sugane, K., 1992, Anisakidae and 

anisakidosis, in Sun, T., ed., Progress in clinical parasitology, v. 3: New York, Springer-Verlag, p. 43–102.

Jackson, G.J., 1975, The “new disease” status of human 

anisakiasis and North American cases: a review: Journal 

of Milk and Food Technology, v. 38, p. 769–773.

Margolis, L., 1977, Public health aspects of “codworm” 

infection: a review: Journal of the Fisheries Research 

Board of Canada, v. 34, p. 887–898.

Myers, B.J., 1975, The nematodes that cause anisakiasis: 

Journal of Milk and Food Technology, v. 38, p. 774–782.

Oshima, T., 1972, Anisakis and anisakiasis in Japan and 

adjacent area in Morishita, K., Komiya, Y., and Matsubayashi, H. eds., Progress of medical parasitology in Japan, 

v. 4: Tokyo, Meguro Parasitological Museum, p. 301–393.

Oshima, T., 1987, Anisakiasis—is the sushi bar guilty?: 

Parasitology Today, v. 3, p. 44–48.

Sakanari, J.A., 1990, Anisakis—from the platter to the 

microfuge: Parasitology Today, v. 6, p. 323–327.

Smith, J.W., 1999, Ascaridoid nematodes and pathology of 

the alimentary tract and its associated organs in vertebrates, including man: a literature review: Helminthological Abstracts, v. 68, p. 49–96.

Smith, J.W., and Wootten, R., 1978, Anisakis and anisakiasis: Advances in Parasitology, v. 16, p. 93–163. 

Biology and transmission of whaleworm, Anisakis simplex

Anderson, R.C., 2000, The superfamily Ascaridoidea, in

Anderson, R.C., ed., Nematode parasites of vertebrates: 

their development and transmission: Oxon, United Kingdom, CABI Publishing, p. 267–314.

Nagasawa, K., 1990, The life cycle of Anisakis simplex: 

a review, in Ishikura, H. and K. Kikuchi, eds., 

Intestinal anisakiasis in Japan: Tokyo, Japan, SpringerVerlag, p. 31–40.

Anisakiosis and Pseudoterranovosis

Nagasawa , K., and Moravec, F., 1995, Larval anisakid 

nematodes of Japanese common squid (Todarodes pacificus) from the Sea of Japan: Journal of Parasitology, v. 81, 

p. 69–75.

Shimazu, T., 1974, Ecology, in Nippon, Suisan Gakkaishi, 

ed., Gyorui to anisakisu: Tokyo, Koseisha-Koseikaku, 

Fisheries and Marine Service Translation Series no. 3417 

(1975). p. 23 – 43 (in Japanese).

Smith, J.W., and Wootten, R., 1978, Anisakis and anisakiasis: Advances in Parasitology, v. 16, p. 93–163. 

Biology and transmission of sealworm, 

Pseudoterranova decipiens, 

particularly from a fishery viewpoint

Anderson, R.C., 2000, The superfamily Ascaridoidea, in

Anderson, R.C., ed., Nematode parasites of vertebrates: 

their development and transmision: Oxon, United Kingdom, CABI Publishing, p. 267–314.

Bowen, W.D. ed., 1990, Population biology of sealworm 

(Pseudoterranova decipiens) in relation to its intermediate 

and seal hosts: Canadian Bulletin of Fisheries and Aquatic 

Sciences, v. 222, 306 p.

Burt, M.D.B. 1994, The sealworm situation, in Scott, M.E., 

and Smith, G., eds., Parasitic and infectious diseases: 

San Diego, Calif., Academic Press, p. 347–362.

Desportes, G., and McClelland, G., eds., 2001, Sealworms 

in the North Atlantic: ecology and population dynamics: 

Tromso, Norway, NAMMCO Scientific Publications, 

v. 3, 171 p.

Hafsteinsson, H., and Rizvi, S.H., 1987, A review of the 

sealworm problem: biology, implications and solutions: 

Journal of Food Production, v. 50, p. 70–84.

Myers, B.J., 1960, On the morphology and life history 

of Phocanema decipiens (Krabbe, 1878) Myers, 1959 

(Nematoda: Anisakidae): Canadian Journal of Zoology, 

v. 38, p. 331–344.
Publishing support was provided by the USGS Columbus Publishing Service Center.

Editing by Elizabeth Enright.

Layout, illustrations and cover by Rosemary Stenback.
For more information concerning this report, contact:

Director 

U.S. Geological Survey

National Wildlife Health Center

6006 Schroeder Road

Madison, WI 53711-6223

jsleeman@usgs.gov

or visit our Web site at:

http://www.nwhc.usgs.gov/
Measures—Anisakiosis and Pseudoterranovosis—Circular 1393

ISSN 1067-084X (print)

ISSN 2330-5703 (online)

http://dx.doi.org/10.3133/cir1393