Circular 1412

National Wildlife Health Center

Baylisascaris Larva Migrans

U.S. Department of the Interior

U.S. Geological Survey
1

234

5

6

Cover artwork: 

Background image: Chicago view with Chase Park in foreground, 

photo by YoChicago.com;

1. Common raccoon, Germany, photo by Carsten Volkwein; 

2. Cardinal with neural larva migrans, photo by Kevin Kazacos 

 and Sam Royer;

3. Young dog found to be infected with B. procyonis, photo by Sam Royer; 

4. Allegheny woodrat in an Indiana cave, photo by Kevin Kazacos; 

5. B. procyonis larva from brain squash of conure, photo by Kevin Kazacos 

 and Sam Royer; 

6. Raccoon latrine on rooftop, photo by William J. Murray. 
Baylisascaris Larva Migrans

By Kevin R. Kazacos

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

USGS National Wildlife Health Center

Circular 1412

U.S. Department of the Interior

U.S. Geological Survey
U.S. Department of the Interior

SALLY JEWELL, Secretary

U.S. Geological Survey

Suzette M. Kimball, Director

U.S. Geological Survey, Reston, Virginia: 2016

For more information on the USGS—the Federal source for science about the Earth, its natural and living 

resources, natural hazards, and the environment—visit http://www.usgs.gov or call 1–888–ASK–USGS.

For an overview of USGS information products, including maps, imagery, and publications, 

visit http://www.usgs.gov/pubprod/.

Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the 

U.S. Government.

Although this information product, for the most part, is in the public domain, it also may contain copyrighted materials 

as noted in the text. Permission to reproduce copyrighted items must be secured from the copyright owner.

Suggested citation:

Kazacos, K.R., 2016, Baylisascaris Larva Migrans: U.S. Geological Survey Circular 1412, 122 p., 3 appendixes, 

http://dx.doi.org/10.3133/cir1412.

Library of Congress Cataloging-in-Publication Data

Names: Kazacos, Kevin R., author.

Title: Baylisascaris larva migrans / by Kevin R. Kazacos.

Description: Reston, Viriginia : United States Department of the Interior,

 U.S. Geological Survey, 2016. | Series: U.S. Geological Survey circular ;

 1412

Identifiers: LCCN 2016009973 | ISBN 9781411340299

Subjects: LCSH: Raccoons as carriers of disease. | Baylisascaris procyonis. |

 Raccoon--Parasites.

Classification: LCC RA641.R15 K38 2016 | DDC 614.5/554--dc23

LC record available at https://lccn.loc.gov/2016009973

ISSN 1067-084X (print)

ISSN 2330-5703 (online)
iii

Foreword

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

“Let both sides seek to invoke the wonders of science instead of its terrors. Together let 

us explore the stars, conquer the deserts, eradicate disease, tap the ocean depths, and 

encourage the arts and commerce.”

John F. Kennedy, from his Presidential inaugural address on January 20, 1961

Disease emergence in wildlife since the late 1900s has been of unprecedented scope relative to geographic areas of occurrence, wildlife species affected, and the variety of pathogens 

involved (Friend, 2006; Daszak and others, 2000). This treatise on Baylisascaris larva migrans 

highlights the recent emergence of a zoonotic disease that over the past 35 years has evolved 

and progressed as both an animal and human disease. Like many new zoonotic diseases in 

humans in recent years, the emergence of baylisascariasis is a result of our densely populated, 

highly mobilized, and environmentally disrupted world. As towns and cities expand, and wildlife 

populations increase, the wild land-urban interface broadens and human associations with wildlife become increasingly frequent. With geographic distance and isolation no longer meaningful 

barriers, the opportunities for once isolated diseases to spread have never been greater. Dealing 

with emerging diseases requires the ability to recognize pathogens when they first appear and 

to act appropriately. Because outbreaks often are evident in the nonhuman components of the 

environment before humans are affected, understanding our environment and associated “sentinel” wildlife is a prerequisite to protecting human health. 

Increasingly, society is recognizing that parasitic zoonoses are an important component of 

emerging global infectious diseases (Daszak and Cunningham, 2002), not only for wildlife but 

for human populations. Because over 50 percent of the pathogens involved with human disease have had their origins in wild animal populations (Daszak and others, 2000; World Health 

Organization, 2004), there is more recognition than ever before of the need to better integrate 

the disciplines of human and animal health to address the phenomenon of infectious disease 

emergence and resurgence. Recognizing the need to document the present status of zoonotic 

diseases, the U.S. Geological Survey (USGS) instituted a series of USGS Circulars on emerging 

zoonotic diseases. This summary of baylisascariasis is the eighth in the series. 

Zoonotic diseases, such as baylisascariasis, are receiving increasing attention as components of 

disease emergence and resurgence (Kazacos, 2001). Baylisascariasis is caused by the roundworm Baylisascaris procyonis and is one of the more recent zoonotic disease developments. This 

disease remains one of the least known and poorly understood zoonotic diseases, yet over the 

past several decades it has become widespread. It originated in wildlife species and is now well 

established as a human malady. Baylisascariasis is transmitted to humans via consumption of 

contaminated feces, and the role of wildlife (primarily raccoons) in this transmission process is 

becoming more clearly known and is outlined in this report. Currently, over 50 percent of the raccoons in the United States are infected, and a smaller percentage in Europe is infected. Because 
iv

raccoons have been relocated from the United States to Europe and Asia, this zoonotic disease 

also causes problems in wildlife species across the globe as well as being a major cause of 

concern for human health. Future generations of humans will continue to be jeopardized by Baylisascaris infections in addition to many of the other zoonotic diseases that have emerged during 

the past century. Through monitoring Baylisascaris infection levels in wildlife populations, we 

will be better able to predict future human infection levels of this important zoonotic disease. 

“In examining disease, we gain wisdom about anatomy and physiology and biology. In 

examining the person with disease, we gain wisdom about life.”

Oliver Sacks 

References Cited

Daszak, P., and Cunningham, A.A., 2002, Emerging infectious diseases: A key role for conservation 

medicine, in Aguirre, A.A., Ostfeld, R.S., House, C.A., Tabor, G.M., and Pearl, M.C., eds., Conservation 

medicine: Ecological health in practice: New York, Oxford University Press, p. 40–61.

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: U.S. Geological 

Survey Circular 1285, 388 p. 

Kazacos, K.R., 2001, Baylisascaris and related species, in Samuel, W.M., Pybus, M.J., and Kocan, A.A., 

eds., Parasitic diseases of wild mammals (2nd ed).: Ames, Iowa State University Press, p. 301–341.

World Health Organization, 2004, Expert consensus, in Contruvo, J.A., Dufour, A., Rees, G., Bartram, J., 

Carr, R., Cliver, D.O., Craun, G.F., Fayer, R., and Gannon, V.P.J., eds., Waterborne zoonoses: Identification, causes and control: London, IWA Publishing, p. 3–16.
v

Contents

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

Acknowledgments ........................................................................................................................................x

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

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

Causative Agent...........................................................................................................................................11

Geographic Distribution..............................................................................................................................14

Patterns and Trends ....................................................................................................................................26

Species Susceptibility.................................................................................................................................31

Human Infections................................................................................................................................31

Animal Infections................................................................................................................................42

Raccoons.....................................................................................................................................42

Paratenic Hosts..........................................................................................................................42

Infecting Dose and Onset of Disease............................................................................57

Clinical Signs .....................................................................................................................58

Granuloma Formation.......................................................................................................61

Obtaining a Diagnosis .................................................................................................................................62

Disease Ecology...........................................................................................................................................66

Points to Ponder...........................................................................................................................................72

Disease Prevention and Control................................................................................................................73

Reducing Environmental Contamination.........................................................................................73

Depopulation and Removal ......................................................................................................73

Anthelmintic Baiting..................................................................................................................76

Preventing Contact with Contaminated Areas or Articles...........................................................76

Education..............................................................................................................................................77

Treatment .............................................................................................................................................77

Intestinal Worms .......................................................................................................................77

Migrating Larvae........................................................................................................................78

Preventive Treatment................................................................................................................78

References Cited..........................................................................................................................................84

Glossary.......................................................................................................................................................102

Appendix 1. Common and scientific names of animals cited .........................................................110

Appendix 2. Cases of Baylisascaris neural larva migrans in pets .................................................117

Appendix 3. Cases of Baylisascaris neural larva migrans in animal facilities.............................118
vi

Figures

1. Photograph showing adult Baylisascaris procyonis roundworms from an 

Indiana raccoon..........................................................................................................................11

2. Photographs showing common raccoon................................................................................14

3. Map showing distribution of the common raccoon in North and Central America ........15

4. Map showing distribution of the common raccoon in Europe............................................15

5. Map showing distribution and general prevalence of Baylisascaris procyonis in 

raccoons in the United States and Canada............................................................................23

6. Map showing numbers and locations of animal cases and outbreaks of 

Baylisascaris larva migrans in the United States and Canada...........................................24

7. Map showing locations of human cases of Baylisascaris larva migrans in the 

United States and Canada.........................................................................................................25

8. Diagram showing the life cycle of Baylisascaris procyonis ...............................................43

9. Photographs showing mammals affected with Baylisascaris procyonis neural 

larva migrans ..............................................................................................................................58

10. Photographs showing the sequence of clinical neural larva migrans in a wild gray 

squirrel..........................................................................................................................................59

11. Photograph showing abnormal circling behavior of a mouse or vole in the snow 

along the Bighorn River near Worland, Wyoming, likely caused by Baylisascaris

neural larva migrans ..................................................................................................................59

12. Photographs showing birds affected with Baylisascaris procyonis neural larva 

migrans.........................................................................................................................................60

13. Photograph showing larval granulomas on the intestinal tract of a hamster with 

B. columnaris neural larva migrans.........................................................................................61

14. Diagram showing areas near homes where humans and animals are at risk of 

contacting raccoon feces containing B. procyonis eggs....................................................66

Topic Highlight Boxes

A History of Baylisascaris Neural Larva Migrans....................................................................................5

A History of Baylisascaris Ocular Larva Migrans ....................................................................................9

Life Cycle of Baylisascaris procyonis ......................................................................................................12

Other Hosts of Adult Baylisascaris procyonis ........................................................................................29

Clinical Disease in Humans........................................................................................................................37

Diagnosis of Baylisascaris Neural Larva Migrans.................................................................................63

Latrines and Eggs—This is the Dangerous Part ....................................................................................67

Plight of the Allegheny Woodrat ...............................................................................................................74

Decontamination Procedures for Baylisascaris Eggs ..........................................................................79

Coping with Baylisascaris Neural Larva Migrans—

From Individual People, to Wildlife Populations, to Organizations ........................................81
vii

Tables

1. Mammals naturally and (or) experimentally affected by Baylisascaris neural larva 

migrans as of 2015 ........................................................................................................................2

2. Birds naturally and (or) experimentally affected by Baylisascaris neural larva 

migrans as of 2015 ........................................................................................................................3

3. Taxonomy of Baylisascaris spp................................................................................................11

4. Baylisascaris species and their primary hosts......................................................................11

5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis

in raccoons ..................................................................................................................................16

6. Location, type of infection, number of animals infected, and numbers of intestinal 

Baylisascaris procyonis, B. potosis, or Baylisascaris spp. in nonraccoon definitive 

hosts..............................................................................................................................................28

7. Primates seropositive for Baylisascaris larva migrans........................................................32

8. Cases of Baylisascaris neural larva migrans in humans.....................................................34

9. Cases of Baylisascaris ocular larva migrans-diffuse unilateral subacute 

neuroretinitis in humans............................................................................................................40

10. Rodents with naturally occurring Baylisascaris neural larva migrans .............................44

11. Rabbits with naturally occurring Baylisascaris neural larva migrans ..............................47

12. Carnivores with naturally occurring Baylisascaris neural larva migrans ........................48

13. Primates with naturally occurring Baylisascaris neural larva migrans............................49

14. Miscellaneous mammals with naturally occurring Baylisascaris neural larva 

migrans.........................................................................................................................................50

15. Gamebirds with naturally occurring Baylisascaris neural larva migrans. .......................50

16. Perching birds with naturally occurring Baylisascaris neural larva migrans..................51

17. Wading birds, shorebirds, diving birds, and waterfowl with naturally occurring 

Baylisascaris neural larva migrans .........................................................................................51

18. Parrots with naturally occurring Baylisascaris neural larva migrans...............................52 

19. Flightless birds with naturally occurring Baylisascaris neural larva migrans.................53

20. Miscellaneous birds with naturally occurring Baylisascaris neural larva migrans .......54

21. Animals experimentally susceptible to Baylisascaris neural larva migrans....................55

22. Clinical signs of Baylisascaris neural larva migrans in small mammals...........................58

23. Clinical signs of Baylisascaris neural larva migrans in birds..............................................60

24. Examples of locations of Baylisascaris larval granulomas in mammals ..........................62
viii

Conversion Factors

International System of Units to Inch/Pound

Multiply By To obtain

Length

inch (in.) 2.54 centimeter (cm)

inch (in.) 25.4 millimeter (mm)

mile (mi) 1.609 kilometer (km)

International System of Units to Inch/Pound

Multiply By To obtain

Length

centimeter (cm) 0.3937 inch (in.)

millimeter (mm) 0.03937 inch (in.)

Area

hectare (ha) 2.471 acre

hectare (ha) 0.003861 square mile (mi2

) 

Mass

gram (g) 0.03527 ounce, avoirdupois (oz)

Temperature in degrees Celsius (°C) may be converted to degrees Fahrenheit (°F) as follows: 

°F=(1.8×°C)+32

Temperature in degrees Fahrenheit (°F) may be converted to degrees Celsius (°C) as follows: 

°C=(°F–32)/1.8

A micrometer (μm) is a unit of measurement of the sizes of cells and bacteria. One micrometer 

is equal in length to one-thousandth of a millimeter.

Words shown in bold are defined in the glossary.
ix

Abbreviations

CDC Centers for Disease Control and Prevention

CNS Central nervous system

CSF Cerebrospinal fluid

DUSN Diffuse unilateral subacute neuroretinitis

ELISA Enzyme-linked immunosorbent assay

LM Larva migrans

MRI Magnetic resonance imaging

NLM Neural larva migrans

NWHC National Wildlife Health Center

OLM Ocular larva migrans

PI Postinfection

VLM Visceral larva migrans

USGS U.S. Geological Survey
x

Acknowledgments

This publication is based on many years of research, investigation, research collaboration, and 

assistance from many individuals spanning a wide variety of professions and disciplines. These 

include numerous veterinarians, physicians, pathologists, diagnosticians, wildlife and exotics biologists, public health officials, parasitology researchers, students, staff, animal owners, 

human patients and their families, and others I have worked with. I thank them all for their 

generosity, cooperation, diligence, and assistance in providing and sharing information, giving 

access to case and reference material, collecting and providing samples, processing specimens, 

and allowing us to learn more about this exceptionally interesting parasite. Without everyone’s 

help, the extent of our progress would not have been possible, and it is sincerely appreciated. 

As with many things in life, my long association with Baylisascaris resulted from serendipity, 

based on a very interesting case that crossed my desk. Who would think that more than 35 years 

later I would still be working on it and that with the help of my numerous colleagues, we would 

gain such a great understanding of the importance of this devastating parasite? My sincere 

thanks to all. 

Special thanks to Rachel C. Abbott for her editorial suggestions and assistance in the development and organization of this publication.

The author with a common raccoon. 

(Photo by Sam Royer)
Baylisascaris Larva Migrans

By Kevin R. Kazacos1

“.....the experiments reported here suggest that 

B. procyonis should be considered dangerous to man 

until proved otherwise.” (Kazacos, Wirtz, and others, 

1981)

“The evidence presented overwhelmingly supports the 

conclusion that this patient died of Baylisascaris eosinophilic meningoencephalitis.” (Huff and others, 1984)

Synonyms

Baylisascariasis, baylisascarosis, raccoon roundworm infection, raccoon ascarid infection, raccoon roundworm encephalitis, Waschbärenspulwurm Infektionen (German)

Overview

Baylisascaris procyonis, the common raccoon roundworm or ascarid, an intestinal parasite, is the most commonly 

recognized cause of clinical larva migrans (LM) in animals, 

a condition in which an immature parasitic worm or larva

migrates in a host animal’s tissues, causing obvious disease, 

but the larva does not develop to the adult stage. Infection 

with B. procyonis is best known as a cause of fatal or severe 

neurologic disease that results when the larvae invade the 

brain, the spinal cord, or both; this condition is known as 

neural larva migrans (NLM) and, historically, cerebrospinal nematodiasis. NLM has been seen in over 150 species

of mammals and birds in North America and elsewhere 

(tables 1 and 2). 

Baylisascariasis is a zoonotic disease, that is, one that is 

transmissible from animals to humans. In humans, Baylisascaris procyonis can cause damaging visceral (VLM), ocular

(OLM), and neural larva migrans (Sorvillo and others, 2002; 

Murray and Kazacos, 2004; Gavin and others, 2005; Wise and 

others, 2005; Shafir and others, 2006; Saffra and others, 2010; 

Kazacos and others, 2013; Singaravelu and others, 2016). Due 

to the ubiquity of infected raccoons around humans, there is 

considerable human exposure and risk of infection with this 

parasite (Roussere and others, 2003; Page, Anchor, and others, 

2009). In North America, Baylisascaris has gained notoriety 

as a cause of eosinophilic meningoencephalitis mainly in 

children, and human cases are being increasingly recognized 

(Murray, 2004; Murray and Kazacos, 2004; Gavin and others, 

2005; Perlman and others, 2010; Kazacos and others, 2013). 

The remarkable disease-producing capability of B. procyonis

in animals and humans is one of the most significant aspects 

of the biology of ascarids (large roundworms) to come to light 

in recent years (Kazacos, 2001). Infection with B. procyonis 

has important health implications for a wide variety of freeranging and captive wildlife, zoo animals, domestic animals, 

as well as human beings, on both an individual and population level (Page, 2013). Other closely related ascarids, for 

example, B. columnaris of skunks and B. melis of badgers, are 

also potential causes of clinical larva migrans in animals and 

humans. 

Background

Baylisascaris procyonis was first reported during the 

1930s in raccoons in the New York Zoological Park (McClure, 

1933) and on a fur farm in Minnesota (Olsen and Fenstermacher, 1938). At the time, the parasite was considered to be 

Ascaris columnaris, which had been previously described in 

skunks. It was later described as a distinct species (Ascaris 

procyonis) based on worms collected from raccoons in Poland 

(Stefański and Žarnowski, 1951). Subsequently, the parasite 

was renamed and included in the newly erected genus

Baylisascaris (Sprent, 1968).

Within the last 60 or so years, and especially the last 35, 

baylisascariasis has evolved and progressed as both an animal 

and human disease. In the late 1940s and early 1950s, Drs. 

Jack Tiner and John F.A. Sprent did considerable research on 

larval migration, larval distribution in mice, growth of the 

larvae, and central nervous system (CNS) disease capability, and compared the raccoon and skunk parasites to other 

carnivore ascarids that had similar somatic migration within 

the body tissues of paratenic (or transport) hosts (Tiner, 

1949, 1951, 1952a,b, 1953a,b; Sprent, 1951, 1952a,b, 1953a,b, 

1955). Their excellent early work set the stage for the clinical importance of Baylisascaris. A few naturally occurring 

cases and outbreaks of clinical NLM were described in the 

1960s and 1970s but, especially in early reports, the parasites involved were not conclusively identified. Concerning 1.

 Professor Emeritus, Purdue University College of Veterinary Medicine
2 Baylisascaris Larva Migrans

Table 1. Mammals naturally and (or) experimentally affected by Baylisascaris neural larva migrans as of 2015.

[N, naturally susceptible; E, experimentally infected]

Rodents (39 species: 32 naturally susceptible, 16 experimentally infected)

House mouse, wild (N, E) House mouse, laboratory mouse (E) White-footed mouse (N, E)

Deer mouse (N) Brush mouse (N) California pocket mouse (N)

Western harvest mouse (E) Meadow jumping mouse (N, E) Meadow vole (E)

Prairie vole (E) Norway or laboratory rat (E) Allegheny woodrat (N; E)

Dusky-footed woodrat (N) Hispid cotton rat (E) Mongolian gerbil (E)

Muskrat (N) Golden hamster (N, E) Eastern chipmunk (N, E)

Eastern gray squirrel (N, E) Eastern fox squirrel (N) Red-tailed squirrel (N)

American red squirrel (N) Prevost’s squirrel (N) Douglas squirrel (N)

Thirteen-lined ground squirrel (N) California ground squirrel (N) Black-tailed prairie dog (N)

Woodchuck (N, E) Western pocket gopher (N) Botta’s pocket gopher (N)

Guinea pig (domesticated) (N, E) Patagonian mara (N) Capybara (N)

Long-tailed chinchilla (N) North American porcupine (and 

western Canadian subspecies; N) Brazilian porcupine (N)

North American beaver (N) Mountain beaver (N) Nutria (N) 

Hares, rabbits, and pikas (3 species: all naturally susceptible, 2 experimentally infected)

Eastern cottontail (N, E) Desert cottontail (N) European rabbit (domesticated; N, E)

Carnivores (7 species: 5 naturally susceptible, 3 experimentally infected)

Domestic dog (N, E) Red (silver) fox (N) American badger (N)

Southern sea otter (N) Long-tailed weasel (N) Least weasel (E)

Domestic ferret (E)

Primates (24 species: 22 naturally susceptible, 3 experimentally infected)

Black-and-white ruffed lemur (N) Red ruffed lemur (N) Ring-tailed lemur (N)

White-headed lemur (N) Coquerel’s giant mouse lemur (N) Mohol bushbaby (N)

White-headed marmoset (N) Black-mantled tamarin (N) Red-handed (Midas) tamarin (N)

Emperor tamarin (N) Golden-headed lion tamarin (N) Cottontop tamarin (N)

Squirrel monkey (E) De Brazza’s monkey (N) Spider monkey (N)

White-eared titi monkey (N) Mantled guereza (N) Japanese macaque (N) 

Rhesus macaque (N) Crab-eating macaque (N, E) Olive baboon (E)

White-handed gibbon (N) Bornean orangutan (and hybrid; N) Human (N)

Marsupials (5 species: all naturally susceptible)

Red kangaroo (N) Yellow-footed rock wallaby (N) Woylie (N)

Long-nosed potoroo (N) Bennett’s wallaby (N)

Bats (2 species: both naturally susceptible)

Indian flying fox (N) Rodrigues flying fox (N)

Ungulates (2 species: both naturally susceptible)

Domestic sheep (N) Domestic cow (N)
Background 3

Table 2. Birds naturally and (or) experimentally affected by Baylisascaris neural larva migrans as of 2015.

[N, naturally susceptible; E, experimentally infected]

Domestic and wild gamebirds (11 species: all naturally susceptible, 1 experimentally infected)

Domestic chicken (N, E) Chukar (N) Ruffed grouse (N)

Common pheasant (N) Wild turkey (N) Indian peafowl (N)

Northern bobwhite (N) California quail (N) Australian brush turkey (N)

Rock partridge (N) Helmeted guineafowl (N)

Perching birds (14 species: all naturally susceptible)

House sparrow (N) House finch (N) Canary (N)

Bushtit (N) Spotted towhee (N) Loggerhead shrike (N)

California thrasher (N) Northern cardinal (N) Northern mockingbird (N)

American robin (N) European starling (N) Blue jay (N)

Western scrub jay (N) American crow (N)

Parrots (29 species: all naturally susceptible, 1 experimentally infected)

Budgerigar (N) Blue-and-yellow macaw (N) Scarlet macaw (N)

Red-and-green macaw (N) Military macaw (N) Eastern rosella (N)

Blue-fronted Amazon (N) Yellow-headed Amazon (N) Yellow-naped Amazon (N)

Cuban Amazon (N) Red-crowned Amazon (N) Thick-billed parrot (N)

Burrowing parrot (N) African grey parrot (N) Blue-crowned parakeet (N)

Orange-fronted parakeet (N) Sun parakeet (N) Rosy-faced lovebird (N)

Rainbow lorikeet (N) Swainson’s lorikeet (N) Yellow-backed lorikeet (N)

Ornate lorikeet (N) Marigold (Edward’s) lorikeet (N) Cockatiel (N, E)

Galah cockatoo (N) Red-tailed black cockatoo (N) Salmon-crested cockatoo (N)

White (umbrella) cockatoo (N) Hybrid of blue-and-yellow 

and scarlet macaw (N)

Pigeons and doves (3 species: all naturally susceptible)

Rock pigeon (N) Mourning dove (N) Diamond dove (N)

Woodpeckers (1 species: naturally susceptible)

Northern flicker (N)

Roadrunners (1 species: naturally susceptible)

Greater roadrunner (N)

Mousebirds (2 species: both naturally susceptible)

Speckled mousebird (N) Blue-naped mousebird (N)

Hornbills and Hoopoe (1 species: naturally susceptible)

Northern red-billed hornbill (N)

Owls (1 species: naturally susceptible)

Barn owl (N)

Wading birds and waterfowl (6 species: 5 naturally susceptible, 1 experimentally infected)

Black-crowned night heron (N) Sanderling (N) Inca tern (N)

Mallard X (N) Domestic duck (E) Crested screamer (N)

Flightless birds (3 species: all naturally susceptible)

Emu (N) Ostrich (N) Greater rhea (N)
4 Baylisascaris Larva Migrans

paratenic hosts of Baylisascaris, such as small mammals and 

birds, these used to be called intermediate hosts (Kazacos, 

1983a, 1986, 2001); however, larval development, specifically a larval molt to a subsequent stage, has never been found 

in them (D.D. Bowman, written commun., 2015), indicating 

that no required or essential development takes place, so they 

instead should be called paratenic hosts.

Additional research at the Purdue University College of 

Veterinary Medicine in the 1980s indicated that B. procyonis was highly pathogenic and dangerous, as it killed nearly 

every animal species and individual that was infected, including primates (Kazacos, 1981; Kazacos, Wirtz, and others, 

1981; Kazacos, Vestre, and Kazacos, 1984; Wirtz, 1981, 1982; 

Kazacos and Wirtz, 1983). Surprisingly, fewer natural cases 

than expected were recognized, given the parasite’s ability 

to cause disease (pathogenicity) and its widespread occurrence in raccoons. However, the diagnosis was most likely 

missed because of a diagnostic focus on rabies. Every year 

in the United States and Canada, thousands of rodents and 

rabbits suffering from nervous system disease were (and still 

are) submitted to diagnostic laboratories or health departments, but they were primarily tested as rabies suspects, not 

for Baylisascaris (Fitzpatrick and others, 2014). As one would 

expect, almost all (99 percent) of these animals were negative for rabies, which begged the question as to what they 

were actually suffering from. Unfortunately, the brain and 

other tissues were then unavailable for further testing. When 

similar animals were examined for Baylisascaris larvae and 

other causes, NLM was found to be a common and widespread 

cause of their neurologic disease (Richter and Kradel, 1964; 

Swerczek and Helmboldt, 1970; Fleming and Caslick, 1978; 

Kazacos, Appel, and Thacker, 1981; Roth and others, 1982). 

These diagnoses and documentation of many other cases (for 

example, Richardson and others, 1980; Reed and others, 1981; 

Koch and Rapp, 1981; Kazacos, 1982; Kazacos and others, 

1982; Kazacos, Winterfield, and Thacker, 1982; Kazacos, 

Reed, and others, 1983, Kazacos, Reed, and Thacker, 1986; 

Kazacos, Fitzgerald, and Reed, 1991; Larson and Greve, 1983; 

Myers and others, 1983; Dixon and others, 1988; Thomas, 

1988; Medway and others, 1989; Sanford, 1991; Kwiecien and 

others, 1993) prompted a reconsideration of diagnostic protocols for such animals in laboratories across North America, 

with Baylisascaris NLM given much stronger consideration. 

In addition, considerable research was initiated on the prevalence and natural ecology of Baylisascaris, including how it is 

maintained and transmitted in nature.

The significant zoonotic potential of B. procyonis was 

recognized early on, and studies of the infection in two species 

of nonhuman primates were conducted (Kazacos, Wirtz, and 

others, 1981; Kazacos, Vestre, and Kazacos, 1984; Kazacos 

and others, 1985). The infecting doses were relatively low 

and easily transmittable to young children, which further put 

the zoonotic risk of B. procyonis into perspective. During 

this time, cases of human OLM and NLM were identified, 

and the NLM cases conclusively linked to B. procyonis from 

raccoons (Raymond and others, 1978; Gass and Braunstein, 

1983; Huff and others, 1984; Fox and others, 1985). As would 

be expected for a previously unknown human disease, other 

earlier cases in humans that matched a probable diagnosis of 

baylisascariasis were identified in the literature, and these are 

now considered to be additional human cases of Baylisascaris

NLM and OLM, including one from Europe (Parsons, 1952; 

Schrott, 1961; Anderson and others, 1975).

Since the 1970s, the parasite has been increasingly identified as a cause of animal and human disease. Over 2,400 cases 

of raccoon roundworm encephalitis in animals and 25 cases 

in humans have been published or reported (Kazacos, 2001; 

Kazacos, Gavin, and others, 2002; Schultz, 2002; Murray and 

Kazacos, 2004; Cheney, 2005; Chris, 2005; Gavin and others, 

2005; Shafir and others, 2006; Pai and others, 2007; Reilly, 

2008; Chun and others, 2009; Hajek and others, 2009; Kelly 

and others, 2009, 2012; Moore, 2009; Mehta and others, 2010; 

Perlman and others, 2010; Ciarlini and others, 2011; Hung and 

others, 2012; Haider and others, 2012; Kazacos and others, 

2013) (table 8), and additional cases of animal and human 

NLM as well as numerous cases of covert or subclinical

infection as determined through serology have also been identified (Cunningham and others, 1994; Brinkman and others, 

2003; K.R. Kazacos, unpub. data, 1995–2010). In addition, the 

parasite has progressed from a questioned cause of human eye 

disease to a well-known cause of human OLM and the primary 

cause of the large nematode variant of the ocular syndrome, 

diffuse unilateral subacute neuroretinitis (DUSN).
A History of Baylisascaris Neural Larva Migrans

Background 5

A. Chickens suffering from Baylisascaris neural 

larva migrans. (Photo by Sam Royer)

The stage was set for Baylisascaris to enter the scene 

as an agent of animal and human disease when two pioneering parasitologists, Dr. Jack D. Tiner and Dr. John F. A. 

Sprent, began examining the biology of larval migration due 

to these parasites in the late 1940s and early 1950s (see references ). It was their research that showed B. procyonis and 

B. columnaris larvae aggressively migrate through somatic 

tissues and tend to cause severe clinical central nervous system disease (neural larva migrans, NLM) in paratenic hosts. 

Dr. Tiner worked primarily with B. procyonis of raccoons 

and Dr. Sprent with B. columnaris of skunks, which previously were considered the same or similar species from the 

two respective hosts. They also worked with other species, 

such as B. melis of badgers, B. transfuga of bears, B. devosi

of weasels, and B. tasmaniensis of Tasmanian devils. Their 

work was meticulous, intriguing, and sometimes downright 

scary. Indeed, a graduate student roommate of Dr. Tiner 

was quite concerned about what Jack was working on, since 

his mice would circle, fall over, and die of nervous system 

disease. And then there were the wild house mice in the lab 

building that also came down with it, apparently having gotten into some of the egg-laden peanut butter used to infect 

the experimental mice. The other students tended to give 

Jack and his animals a wide berth.

Tiner’s and Sprent’s work showed differences in the 

pathogenicities of the raccoon and skunk ascarids and a 

greater disparity between these two species and the bear, 

weasel, dog, and cat ascarids as causes of clinical NLM. 

They found that Baylisascaris of raccoons and skunks are 

aggressive migrators whose larvae enter the tissues of mice 

and other rodents soon after oral infection with infective 

eggs. As part of this migration, small numbers (approximately 5–7 percent) of larvae enter the brain, but because 

of their rapid growth and aggressiveness, they produce considerable damage to the brain, resulting in clinical disease 

(Tiner, 1953b; Sprent, 1955). The raccoon ascarid was about 

20 times more pathogenic than the skunk ascarid, whose larvae were walled off more readily by the host and thus were 

better tolerated; however, the skunk roundworm could still 

produce similar brain disease in some mice. Tiner (1953a,b) 

also found that the badger roundworm (B. melis) was a 

particularly pathogenic species in causing NLM, and Sprent 

(1952a, 1953a,b, 1955) showed that the ascarids of bears, 

weasels, and dogs were not nearly as pathogenic as the others. Based on the experimental work of Tiner and Sprent, 

another famous parasitologist, Dr. Paul Beaver, first alluded 

to the zoonotic potential of Baylisascaris and included these 

parasites in his discussion of larva migrans and paratenesis 

(Beaver, 1969). In addition to studying migration and neural 

pathogenicity, Tiner first noted the relevance of B. procyonis as a cause of natural disease in animal populations and 

studied the parasite in squirrels in Illinois woodlots. Tiner 

estimated that B. procyonis was responsible for as much 

as 5 percent of natural rodent mortality in woodlots where 

raccoons were common (Tiner, 1954). Despite the excellent early research of these two scientists, many still viewed 

these parasites as interesting curiosities, and the widespread 

importance of Baylisascaris as disease agents would remain 

hidden until later.

Three cases or outbreaks of naturally occurring clinical NLM due to Baylisascaris were described in the 1960s 

and a dozen more in the 1970s, in woodchucks, squirrels, 

long-tailed chinchillas, beavers, nutria, rabbits, and several 

avian species (Richter and Kradel, 1964; Kelly and Innes, 

1966; Fritz and others, 1968; Swerczek and Helmbolt, 1970; 

Schueler, 1973; Church and others, 1975; Dade and others, 

1975, 1977; Nettles and others, 1975; Helfer and Dickinson, 

1976; Jacobson and others, 1976; Fleming and Caslick, 

1978; Sass and Gorgacz, 1978; Winterfield and Thacker, 

1978; Fleming and others, 1979). Again, they were viewed 

for the most part as pathogenic curiosities, and in early 

cases, the identification of the parasite and the source of the 

infection were both lacking. Sprent (1968) had described the 

new genus Baylisascaris and listed the known species, but 

it took some time for this to become widely known. Later 

papers indicated more strongly the broader importance of 

the parasites in causing larger clinical outbreaks (Dade and 

others, 1975, 1977; Nettles and others, 1975; Jacobson and 

others, 1976). 

In 1979, the owner of a poultry operation in central 

Indiana was steadily losing birds to rapidly progressive 

central nervous system (CNS) disease: 115 birds in the 

first week, 66 in the second, 109 in the third, etc. 

(A). Birds were submitted to the Indiana Animal Disease 

Diagnostic Laboratory, but the cause was undetermined. 
A History of Baylisascaris Neural Larva Migrans

6 Baylisascaris Larva Migrans

Viral encephalitis was a possibility because the birds had 

not been vaccinated against Newcastle disease. However, 

histological examination of brain tissue revealed lesions that 

looked like something had migrated through the brain. Upon 

examining sequential thin sections of the brains of several 

birds, multicellular parasites were found in the brain tissue 

and were identified as large ascarid larvae (B).

Living larvae were isolated from the brains of other 

birds by using a Baermann funnel (C). The parasites 

were identified as larvae of Baylisascaris, most likely 

B. procyonis. 

One and a half years later, similar lesions and parasites were seen in bobwhites with neurologic disease (F). 

The bobwhites were from a group of 85 that had access to 

a 12-by-24 foot dirt pen in which three baby raccoons had 

been housed previously for three months (G). In short order, 

all of the birds died from CNS disease (Reed and others, 

1981). Subsequent investigation indicated that even though 

the raccoons were lightly infected with Baylisascaris, they 

were shedding 1,300–5,400 eggs per gram of feces and had 

contaminated the pen with an estimated 155,593,800 eggs 

(Kazacos, 1982), ensuring the death of all of the birds from 

NLM. Indeed, Tiner and Sprent had shown that a single 

larva in the brain of a small rodent was fatal.

Within two years of the original outbreak, several 

additional cases were identified: a woodchuck submitted as 

a rabies suspect, a brushturkey from the Indianapolis Zoo, 

an emu (with B. columnaris), and domestic rabbits suffering 

high mortality (again, contaminated straw was the source) 

(Kazacos, Appel, and Thacker, 1981; Kazacos, Kazacos, 

and others, 1982; Kazacos, Winterfield, and Thacker, 1982; 

Kazacos and others, 1983). Mice, hamsters, rats, gray squirrels, chickens, ducks, ferrets, a chipmunk, woodchucks, 

least weasel, squirrel monkeys, and crab-eating macaques 

were experimentally infected with B. procyonis eggs (H, I), 

and nearly all died with severe CNS disease (Kazacos, 1981, 

1983a; Wirtz, 1982; Kazacos and Wirtz, 1983; Kazacos, 

Wirtz, and others, 1981; Kazacos, Vestre, Kazacos, and Raymond, 1984; Kazacos, Vestre, and Kazacos, 1984; Kazacos 

and others, 1985; Kazacos and Kazacos, 1984, 

Kazacos, E.A., and Kazacos, K.R.,1988).

B. Histology of cross-sections 

of Baylisascaris larvae in 

chicken brain. (Photo by Kevin 

Kazacos and Sam Royer)

C. Baylisascaris larva recovered 

from brain of chicken. (Photo by 

Kevin Kazacos and Sam Royer)

D. Raccoon trapped 

from barn. (Photo by 

Kevin Kazacos)

E. Raccoon feces 

recovered from straw 

loft of barn. (Photo by 

Kevin Kazacos and 

Sam Royer)

The owner continued to lose birds, although fatalities 

were decreasing by the sixth week. The birds were housed 

in a total confinement building, with little human and 

no animal traffic into the poultry house. The owner was 

presented with a list of carnivores (dogs, cats, raccoons, 

skunks, weasels, foxes, coyotes, and bears), and asked if 

there might be any contact with any of them. He immediately picked out raccoons, and said he had a terrible problem with them denning and defecating in his straw loft 

(D, E). This was the same straw he had unwittingly used 

as litter in his poultry house, revealing how the birds were 

becoming infected. B. procyonis eggs were identified in 

the raccoon feces from the straw loft, and the owner was 

advised to 1) not use any more of the contaminated straw, 

2) remove the birds and carefully clean out all of the material from the poultry house and straw loft and burn it, and 

3) prevent any further raccoon incursions and contamination. He did this right away; losses essentially ceased 

by the end of the seventh week, and subsequent groups 

of birds did not become infected. In total, the owner lost 

622 birds over the 7-week period, and until recently this 

remained the largest outbreak of Baylisascaris encephalitis ever recorded (Richardson and others, 1980). 
A History of Baylisascaris Neural Larva Migrans

Background 7

I. Hamsters with Baylisascaris neural larva migrans, ten 

days after experimental infection with B. procyonis eggs. 

(Photo by Kevin Kazacos and Sam Royer)

Research using squirrel monkeys and crab-eating 

macaques provided important insights into the pathogenicity 

of the parasite in primates and thus potentially in humans 

(Kazacos, Wirtz, and others, 1981; Kazacos, Vestre, and 

Kazacos, 1984; Kazacos and others, 1985). All of these 

primates died from CNS disease, and the parasites had 

migrated extensively throughout muscle and connective 

tissue, the brain, and the eyes, proving the probable danger 

of B. procyonis to humans. In addition to data on NLM, the 

macaques provided great insight into clinical Baylisascaris

ocular larva migrans (OLM).

In early 1983, a child in Pennsylvania died of eosinophilic meningoencephalitis and larvae in his tissues were 

identified as Baylisascaris, for the first time ever in a human 

(Huff and others, 1984) (J). Cases of human OLM and diffuse unilateral subacute neuroretinitis (DUSN) were also 

attributed to Baylisascaris (Raymond and others, 1978; Gass 

and Braunstein, 1983; Kazacos, Vestre, Kazacos, and Raymond, 1984; Kazacos, Vestre, and Kazacos, 1984; Kazacos 

and others, 1985). Based on rapidly accumulating evidence, 

H. Mice with Baylisascaris neural larva migrans, ten days 

after experimental infection with B. procyonis eggs. 

(Photo by Kevin Kazacos and Sam Royer)

F. Bobwhite with Baylisascaris neural larva 

migrans. (Photo by Kevin Kazacos)

G. Pet raccoon linked to outbreak of 

Baylisascaris neural larva migrans 

in bobwhites. (Photo by 

Kevin Kazacos)

B. procyonis was described as a cause of both animal and 

human disease (Kazacos, 1983a), and shortly thereafter the 

parasite was identified as the cause of death of a second 

child, in Illinois (K, L) (Fox and others, 1985). All state and 

territorial public health agencies and laboratory directors 

in the country were strongly warned in a letter about B. 

procyonis and its probable zoonotic importance in human 

beings (Kazacos, 1984). In a short time, B. procyonis had 

been identified conclusively for the first time in humans, 

identified in additional cases of NLM, implicated as a cause 

of human OLM and DUSN, and received much deserved 

national attention as a newly recognized human pathogen. 

Based mainly on Baylisascaris, but also including other 

parasites, neural larva migrans was introduced as a new 

type of larva migrans in 1994–95 (Cunningham and others, 

1994; Kazacos, 1995; Kazacos and Boyce, 1989, 1995) and 

has been well accepted. Baylisascaris is now recognized 

internationally as a cause of animal and human disease and 
A History of Baylisascaris Neural Larva Migrans

8 Baylisascaris Larva Migrans

is routinely considered in cases of eosinophilic meningoencephalitis and OLM–DUSN in humans in North America, 

Europe, and Asia. Twenty-five cases of Baylisascaris NLM 

in humans have been published or reported (table 8), over a 

dozen additional cases are also known, and over a hundred 

seropositive individuals have been identified (table 7) (K.R. 

Kazacos, unpub. data, 1995–2010; Brinkman and others, 

2003; Dangoudoubiyam and Kazacos, 2009; Dangoudoubiyam and others, 2011; Rascoe and others, 2013). Excellent serologic tests have been developed for this infection 

and are in current use in at least two national laboratories 

(United States and Canada). Now when infection with Baylisascaris is suspected, serologic confirmation of infection 

is usually sought without delay and aggressive treatment is 

initiated promptly. This has and will continue to make an 

important difference in the lives of those affected by this 

parasite.

K. Areas of dead tissue in the brain of child with neural larva 

migrans in Illinois. (Photo by Nevenka S. Gould)

L. Histology of cross section of Baylisascaris larva in the brain 

of child with neural larva migrans in Illinois. (Photo by 

Nevenka S. Gould)

J. Histology of Baylisascaris larva in the brain of child with 

neural larva migrans in Pennsylvania. (Photo by Kevin Kazacos; 

specimen courtesy of Dale S. Huff)
A History of Baylisascaris Ocular Larva Migrans

Background 9

As a cause of human ocular larva migrans (OLM), 

Baylisascaris was quick to arrive on the scene but slower 

to be accepted. Toxocara spp. of dogs and cats were 

well-known causes of human OLM, having first been 

documented in human eyes in 1950 by pathologist Helenor Wilder (Wilder, 1950) and later identified as Toxocara

(Nichols, 1956). Toxocara larvae do not grow as they 

migrate, remaining small (approximately 400 micrometers 

[μm] long by 16–18 μm in diameter), whereas Baylisascaris

larvae grow quickly, reaching a size of 1,500–2,000 μm 

long by 60–80 μm in diameter in about 2 weeks (Goldberg 

and others, 1993). In studies of B. procyonis in nonhuman 

primates, mainly examining NLM, ophthalmoscopic examinations also were done (A), and it was found that larvae 

entered the eyes of every animal, producing lesions of OLM 

(Kazacos, Wirtz, and others, 1981; Kazacos, Vestre, Kazacos, and Raymond, 1984; Kazacos, Vestre, and Kazacos, 

1984) (B, C). Previously, nematodes in the exact size range 

of Baylisascaris (1,500–2,000 μm) were found in the eyes 

of a 13-year-old girl in Kentucky (D) and a 23-year-old 

man in Michigan (Raymond and others, 1978). The girl had 

obtained a pet raccoon 6 weeks prior to developing visual 

symptoms, suggesting these were previously unrecognized 

cases of Baylisascaris OLM.

In 1983, evidence was presented that a previously 

described ocular syndrome, called diffuse unilateral subacute neuroretinitis (DUSN) (Gass and others, 1978), was 

caused by at least two unidentified nematodes of different 

sizes (Gass and Braunstein, 1983). In 12 patients primarily 

from the southeastern United States, the parasite measured 

400–1,000 μm, and in 6 patients from the Upper Midwest, 

including the 2 patients mentioned above (Raymond and 

others, 1978), the parasite measured 1,500–2,000 μm. B. 

procyonis was suggested as the probable cause of the large 

nematode variant of DUSN, based on the compatible size 

of the larvae, similar disease development by B. procyonis

in animals, including nonhuman primates, the geographic 

location of cases, and the fact that one of the patients had 

acquired a pet raccoon 6 weeks prior to developing ocular 

disease (Kazacos, Vestre, Kazacos, and Raymond, 1984; 

Kazacos, Vestre, and Kazacos, 1984; Kazacos, Raymond, 

and others, 1985).

D. Baylisascaris larva 

(arrow) in the eye 

of a Kentucky girl. 

(Photo by Lawrence 

A. Raymond)

A. Kevin Kazacos (right) and Dr. W.A. Vestre performing an 

ophthalmoscopic exam on a macaque with NLM and 

OLM–DUSN. (Photo by Sam Royer)

B. The interior of 

a macaque’s eye 

showing larva (arrow) 

and migration tracks 

(white areas). (Photo 

by William A. Vestre)

C. Histology of 

larva migrating in 

retina of macaque 

with OLM. (Photo by 

Kevin Kazacos)
A History of Baylisascaris Ocular Larva Migrans

10 Baylisascaris Larva Migrans

During the 1980s, however, this conclusion was controversial to some members of the ophthalmology community. 

Opponents argued that “several patients with DUSN lived in 

highly urban centers in the Northeast and Midwest, making exposure to raccoons highly unlikely” (Lewis, 1985). In 

addition, “no reported patient with DUSN has had neurologic complaints or has presented with meningoencephalitis, as occurs in both natural and experimental infestations 

with B. procyonis in all nonhost species,” and “a number 

of patients with DUSN have had a creeping skin eruption 

during their illness compatible with Ankylostoma [sic]......

cutaneous larva migrans is not a feature of B. procyonis

infestation” (Lewis, 1985). None of these arguments made 

practical or scientific sense for discounting the involvement 

of Baylisascaris in DUSN, because:

1. raccoons are surprisingly common in major urban centers where they have been linked to both human NLM 

and OLM–DUSN (Perlman and others, 2010; Saffra 

and others, 2010; Liu and others, 2015; and table 5); 

2. B. procyonis can cause a full range of clinical problems, from asymptomatic or covert infection at lower 

dosages to central nervous system (CNS) disease at 

higher dosages (Kazacos, 2000; Gavin and others, 

2005; Kazacos and others, 2013); 

3. Baylisascaris OLM would be similar to Toxocara

OLM: primarily, a stand-alone disease resulting from 

low-level infection, with no “requirement” for concomitant NLM or CNS disease (Kazacos, Vestre, and 

Kazacos, 1984; Saffra and others, 2010); and 

4. larva migrans affecting the skin was described for 

the small nematode variant of DUSN, not the large 

(Lewis, 1985; Gass, 1997).

Other ophthalmologists strongly supported B. procyonis as a cause of DUSN and shared additional OLM and 

DUSN cases they had seen that were probably due to Baylisascaris based on exposure history and the larvae that were 

seen. One involved a 42-year-old man in central Wisconsin 

who was an avid outdoorsman and had contact with a likely 

contaminated area (Williams and others, 1988), and another 

involved a 27-year-old man in Pennsylvania from an area 

where raccoons were common (Sivalingam and others, 

1991). There was excellent morphometric, serologic, and 

epidemiologic support for Baylisascaris as the cause of 

OLM–DUSN in a 29-year-old man in northern California 

(Goldberg and others, 1993) (E). A 48-year-old German 

woman who obtained a pet raccoon also developed OLM–

DUSN (Kuchle and others, 1993).

These patients all had large intraocular larvae whose 

size and other characteristics matched Baylisascaris. 

Among the expanding group of patients, several cases of 

Baylisascaris NLM and OLM were reported from urban 

centers, including seropositive cases of Baylisascaris

NLM and OLM in an infant in Brooklyn and a teenager in 

New York City, one of the largest urban centers (Perlman 

and others, 2010; Saffra and others, 2010), Baylisascaris OLM–DUSN in a boy in Chicago and a man in Los 

Angeles (Mets and others, 2003; D.M. Hirota and others, 

unpub. data, 2009), NLM cases from Chicago, Los Angeles and Toronto (Gavin and others, 2002; Kazacos and 

others, 2002; Hajek and others, 2009) and another DUSN 

case in the Bronx in New York City in which the patient 

was positive for Baylisascaris antibodies in her ocular 

fluid (Liu and others, 2015). To date, there are 28 reported 

cases of OLM–DUSN in which the size of the larva, serologic testing, and (or) history implicate Baylisascaris as 

the likely cause (table 9), and many others may exist since 

not all cases are referred or reported. Additional experimental studies have supported the role of this parasite 

in OLM–DUSN (Akao and others, 2003). Baylisascaris

is now well accepted as a cause of human OLM and the 

primary cause of the large nematode variant of DUSN.

E. Baylisascaris larva (arrow) in the eye of California man, also showing 

first attempt at laser treatment to kill larva. (Photo by Barrett Katz)
Causative Agent 11

Causative Agent

Baylisascaris procyonis and its relatives are large roundworms (nematodes, ascarids) (table 3) that live in the small 

intestine of their respective definitive hosts (Kazacos, 2001) 

(fig. 1). Species of Baylisascaris occur primarily in carnivores; 

one species (B. laevis) occurs in rodents (Sprent, 1968; Wu 

and others, 1987; Tokiwa and others, 2014) (table 4). Similar 

to other ascarids of terrestrial carnivores, most Baylisascaris

species are transmitted to definitive hosts when they ingest 

larvae present in small mammal paratenic hosts. Direct infection by ingestion of infective eggs is also important for some 

species, particularly in young definitive hosts (Tiner, 1952a, 

1953a; Sprent, 1953b; Sprent and others, 1973; Kazacos, 

1983b, 2001; Kazacos and Boyce, 1989).

The different species of Baylisascaris vary in how they 

affect the central nervous system of paratenic hosts, and this is 

related to differences in how they migrate through the host’s 

tissues and invade the brain, the growth and aggressiveness 

of larvae in the CNS, and the ability of the host to wall off or 

encapsulate the larvae. Baylisascaris procyonis and B. melis

are the most pathogenic, with aggressive somatic migration 

and brain invasion, followed by B. columnaris and the others 

(Boyce and others, 1988b; Kazacos and Boyce, 1989; Kazacos, 2001).

Experimentally, B. procyonis is biologically different and 

more pathogenic than B. columnaris, requiring host ingestion 

of fewer infective eggs and fewer larvae in the brain to cause 

similar or worse clinical disease. One B. procyonis larva in the 

brain of a mouse is usually fatal, whereas five to six or more 

B. columnaris larvae in the brain are not necessarily fatal, 

even when CNS signs are present (Sprent, 1952a, 1955; Tiner, 

1953a,b; Clark and others, 1969; Sheppard, 1995, 1996; Sheppard and Kazacos, 1997). Clinical signs appear much earlier 

and progress much more quickly in mice infected with B. 

procyonis than with B. columnaris. In addition, B. columnaris

larvae in the brain have a greater tendency to settle down and 

become encapsulated than do larvae of B. procyonis and may 

not produce clinical signs (Tiner, 1953a,b; Sprent, 1955; Clark 

and others, 1969). However, it is also known that, at sufficient 

dosages, B. columnaris has the ability to produce clinically 

significant NLM in susceptible species, including rodents, rabbits, ratites, and nonhuman primates. People should therefore 

exercise the same precautions with B. columnaris as those for 

B. procyonis and B. melis.

Figure 1. Adult Baylisascaris procyonis roundworms from an 

Indiana raccoon. (Photo by Kevin Kazacos and Sam Royer)

Table 3. Taxonomy of Baylisascaris spp.

Classification Designation

Kingdom Animalia

Phylum Nematoda

Class Secernentea

Order Ascaridida

Family Ascarididae

Genus Baylisascaris

Species procyonis

potosis

columnaris

melis

devosi

tranfuga

schroederi

ailuri

tasmaniensis

laevis

Table 4. Baylisascaris species and their primary hosts.

[Compiled from Kazacos (2001); Sprent (1968, 1970); Tokiwa and others 

(2014); Wu and others (1987)]

Species Primary definitive host(s)

B. procyonis Raccoons.

B. potosis Kinkajous.

B. columnaris Skunks.

B. melis Badgers.

B. devosi Martens, fishers.

B. transfuga Bears.

B. schroederi Giant pandas.

B. ailuri Red pandas.

B. tasmaniensis Tasmanian devils, quolls.

B. laevis Marmots, ground squirrels.
Life Cycle of Baylisascaris procyonis

12 Baylisascaris Larva Migrans

1. B. procyonis adults are large, tan-colored worms. Mature females are up to 20–22 centimeters (cm) long, males up to 

9–11 cm long (Sprent, 1968; Gey, 1998; Kazacos, 2001). Both live in the small intestine of raccoons. Adult female 

worms are very prolific and produce an estimated 115,000–179,000 eggs per worm per day; thus, infected raccoons 

shed millions of eggs per day in their feces (Kazacos, 1982, 1983a, 2001; Snyder and Fitzgerald, 1987; Reed and 

others, 2012).

2. The eggs of B. procyonis are roughly oval, brown, contain a large single-celled embryo, and have a thick shell with a 

finely granular or particulate surface. They range in size from 63–88 by 50–70 micrometers (μm), and most average 

68–76 by 55–61 μm (Kazacos and Turek, 1983; Kazacos and Boyce, 1989; Sakla and others, 1989; Miyashita, 1993; 

Averbeck and others, 1995; Van Andel and others, 1995; Conboy, 1996; Gey, 1998).

Adult Baylisascaris

procyonis in raccoon. 

(Photo by Kevin Kazacos)

Baylisascaris adults 

recovered from raccoons 

at necropsy. (Photo by 

Kevin Kazacos) 

Undeveloped 

Baylisascaris procyonis

eggs from fresh raccoon 

feces. (Photo by Kevin 

Kazacos)

Infective B. procyonis egg 

from older raccoon feces. 

Note larva coiled inside. 

(Photo by Kevin Kazacos)

Higher magnification of 

undeveloped 

B. procyonis egg from 

fresh raccoon feces. 

(Photo by Kevin Kazacos)

Early migratory hemorrhages in the 

lungs of a rat and ferret. (Photos by 

Kevin Kazacos and Sam Royer)

3. Given adequate temperature and moisture, B. procyonis

eggs can reach infectivity (third-stage larva) in 11–14 

days (Sakla and others, 1989), although under natural 

conditions of fluctuating temperatures, this usually 

takes several weeks. Once infective, they can remain 

viable in the environment for years (Kazacos and 

Boyce, 1989).

4. Following ingestion by paratenic hosts, infective eggs hatch in the small intestine, and larvae quickly penetrate the intestinal wall and migrate through the liver to the lungs. B. procyonis larvae cause hemorrhages in the lungs of laboratory mice 

and other animals as early as 1 day postinfection (PI), peaking on days 2–3 PI, followed shortly by lung inflammation 

(Wyand-Ouellette and others, 1983; Kazacos, 1986, 2001).
Causative Agent 13

Early migratory 

hemorrhages in the 

brain of a mouse. 

(Photo by Kevin Kazacos 

and Sam Royer)

5. In the lungs, the larvae gain access to the systemic arterial circulation, which distributes them throughout the body but 

especially to the forequarters and head, where some end up in the brain (Sprent 1952a, 1955; Tiner 1953b; Sheppard 

and Kazacos 1997). Some larvae also migrate from the intestine or lungs directly into surrounding tissues.

Histology of Baylisascaris

larva in granuloma in heart of 

a rabbit with NLM. (Photo by 

Kevin Kazacos and 

Sam Royer.)

Baylisascaris procyonis

larva recovered by digestion 

from the brain of a chinchilla. 

(Photo by Kevin Kazacos)

Baby raccoons with Kevin 

Kazacos. (Photo by Sam Royer)

 White-footed mouse, a common 

paratenic host of B. procyonis.

(Photo by Claudia Sheppard and 

Sam Royer)

Larval granulomas on 

heart of a rabbit with NLM. 

(Photo by Kevin Kazacos 

and Sam Royer.)

B. procyonis larva in 

brain squash of a parrot. 

(Photo by Kevin Kazacos 

and Sam Royer)

Life Cycle of Baylisascaris procyonis

6. B. procyonis larvae grow quickly, from about 300 μm long when they 

hatch from the eggs to 1.3–1.9 millimeters (mm) long at 15 days PI 

(Tiner 1953a; Bowman 1987; Donnelly and others, 1989; Goldberg and 

others, 1993). Most B. procyonis larvae recovered from clinical cases 

of neural larva migrans (NLM) are 1.5–1.9 mm long and 60–80 μm in 

greatest diameter (Kazacos, 1997, 2001).

7. Larvae in visceral and somatic tissues become encapsulated in granulomas (inflammatory nodules composed of 

connective tissue, white blood cells, and other cells) where they become quiescent and survive for months to years, 

essentially waiting to be ingested by a raccoon (Kazacos, 1983a, 1997).

8. Young raccoons become infected at an early age by ingesting infective eggs from their 

mother’s contaminated teats or fur, the contaminated den, or while visiting or investigating 

raccoon latrines or other contaminated areas near their den (Kazacos, 1983a,b, 2001).

9. Older raccoons become infected mainly by ingesting third-stage larvae in paratenic 

hosts, usually rodents (Tiner, 1953a,b; Kazacos, 1983a,b, 2001; Kazacos and Boyce, 

1989; Reed and others, 2012), although some egg-induced infections may also occur.

10. In young raccoons, larvae hatching from eggs enter the mucous membranes (mucosa) of the small intestine and develop 

there for several weeks before reentering the intestinal lumen to mature; these worms reach patency (egg shedding) in 

50–76 (mean, 63) days. In older raccoons, larvae from paratenic hosts develop to adult worms in the lumen (interior 

cavity) of the intestine, reaching patency in 32–38 (mean, 35) days (Kazacos, 1983b; Kazacos and Boyce, 1989).
A

B

14 Baylisascaris Larva Migrans

Baylisascaris transfuga, B. devosi, and B. tasmaniensis

are also potential causes of larva migrans in humans; however, 

due to differences in migration, larval growth, and encapsulation of their larvae, they are much less pathogenic than the 

other species. Their larvae mainly enter visceral organs and 

muscle and have much less CNS invasion (Sprent, 1953b; 

Matoff and Komandarev, 1965; Sprent and others, 1973; 

Kazacos and Boyce, 1989; Kazacos, 2001; Papini and others, 1994; Papini, Demi, and others, 1996), although at high 

dosages in mice, both NLM and OLM were produced following infection with B. transfuga (Papini and Casarosa, 1994; 

Papini, Renzoni, and others, 1996; Sato and others, 2004). 

Recently, B. potosis was found to be much less pathogenic 

than B. procyonis in experimentally infected squirrel monkeys. 

At exceptionally high dosages, some larvae became encapsulated at the brain surface, but there was no deep CNS invasion 

or migration and no clinical disease was produced (Tokiwa 

and others, 2015). Larvae mainly migrated in the viscera and 

became encapsulated in the wall of the lower intestine. Less is 

known about the other species; B. schroederi is unusual in that 

it appears to cause clinical LM in its definitive host, the giant 

panda (Zhang and others, 2007; Wang and others, 2008).

The vast majority (greater than or equal to 96 percent) of 

recognized clinical cases of NLM and OLM due to Baylisascaris have been linked to B. procyonis from raccoons (Kazacos, 2001), indicating the much greater importance of this 

species in the causation and epidemiology of these diseases. 

However, all of the above Baylisascaris species, and perhaps 

some others, should be considered potentially dangerous for 

producing larva migrans disease, particularly at higher infection levels.

Geographic Distribution

The geographic distribution of B. procyonis is linked to 

that of the most common host of the adult worm, namely the 

raccoon. Common or North American raccoons (fig. 2) are 

native to North and Central America (Lotze and Anderson, 

1979), and their range and population densities have been 

increasing since the 1940s (Gehrt, 2003) (fig. 3). They have 

also been introduced elsewhere, such that wild populations 

now exist in parts of Europe (fig. 4) and Asia. They are common as exotic pets or in zoological parks elsewhere, including 

in Europe, Japan, and recently described in China (Xie and 

others, 2014). This expansion of raccoons has been accompanied by the spread of B. procyonis to other areas of the world, 

so that B. procyonis is now enzootic in raccoons in North 

America, Europe, and parts of Asia, leading directly to cases 

and outbreaks of neural and ocular larva migrans in animals 

and humans in these areas.

In North America, B. procyonis is more common in the 

Midwestern and Northeastern United States and contiguous 

Canada and on the West Coast, where infection prevalence 

among raccoons reaches 68–100 percent (Kazacos and Boyce, 

1989; Kazacos, 2001) (table 5). The prevalence of B. procyonis appears to be stable in enzootic areas, although some 

differences and fluctuations are possible based on changes 

in the number of animals and their populations (Page, Gehrt, 

and others, 2009), as well as ecoregion and soil characteristics 

(Kresta and others, 2010). In Wisconsin, 75 percent of 

B. procyonis infections are in the southern half of the state 

and 18 percent are in the northern quarter, related to the 

higher abundance of raccoons in the southern part of the state 

versus the northern part (Amundson and Marcquenski, 1986). 

Similar associations of B. procyonis with raccoon abundance 

were seen in Texas (Kresta and others, 2010). In general, the 

prevalence of B. procyonis decreases from the northern to 

southern United States, so that the parasite is less common 

in raccoons in more southern states (Kazacos, 2001) (table 5; 

fig. 5), although pockets of higher prevalence exist there (Kerr 

and others, 1997). 

Figure 2. Common raccoons (A, photo by B.S. Thurner Hof; 

B, photo by Carsten Volkwein). 
0 500 1,000 MILES 250

©2015 BYU Geography

CANADA

MEXICO

UNITED STATES

EXPLANATION

Raccoon distribution area

EXPLANATION

FROM: https://geography.byu.edu/pages/resources/outlinemaps.aspx

BYU Geography has produced the following set of basic outline maps of the world 

Spain

France

Norway

Sweden

Estonia

Russia

Georgia Azerbaijan

Lithuania

Belarus

Ukraine

Romania

Serbia

Germany Poland

Austria

Croatia

Slovakia

Czech. 

Rep.

Hungary

Italy

United

Kingdom

©2015 BYU Geography

0 250 500 Miles

Denmark

Switzerland

Slovenia

Belgium

Luxembourg

Country with raccoons, 

 including sporadic detections

Netherlands

Causative Agent 15

Figure 3. Distribution of the common raccoon in North and Central America. (Modified from Goldman, 1950, 

and Boggess, 1994, and including Pipas and others, 2014.)

Figure 4. Distribution of the common raccoon in Europe. (Modified from Beltrán-Beck and others, 2012)
16 Baylisascaris Larva Migrans

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

North America

Midwestern United States

Midwestern U.S.

(Illinois, Indiana, Kansas, Michigan, 

Minnesota, Missouri, Wisconsin)

1,050 36 6

(1–199)

Page and others (2015).

Illinois 6 4 of 6 27

(2–71)

Leigh (1940).

Central Illinois 1 1 of 1 ~120 Tiner (1952a).

Central Illinois and Wisconsin NR NR NR Tiner (1953a).

Southern Illinois 36 64 NR Barnstable and Dyer (1974).

Northeast Illinois (Chicago) 26 42 21

(1–101)

Pigage and others (1983).

Northeast Illinois (Chicago) 307 39 13 Page and others (2009b).

Northeast Illinois (Chicago) 1

433 18 NR Page and others (2009b).

Illinois 310 82 52

(1–328)

Snyder and Fitzgerald (1985).

Illinois 100 86 52

(1–241)

Snyder and Fitzgerald (1987).

Southern Illinois 60 5 NR Birch and others (1994).

West-central Illinois 1

392 38 NR Mitchell and others (1998).

Indiana 25 28 NR Robinson and others (1957).

Central Indiana 4 3 of 4 NR Reed and others (1981).

Indiana 1

95 20 NR Jacobson and others (1982).

Indiana 2

218 29 NR Jacobson and others (1982).

Indiana 1,425 72 NR Kazacos and Boyce (1989).

1982 sample 391 74 43

(1–283)

K.R. Kazacos, unpub. data 

(1982).

Northwest Indiana 2

219 15 NR Cooney (1989).

Iowa 1 1 of 1 Quite

abundant

Morgan and Waller (1940).

Iowa 10 20 NR Waller (1940).

Iowa 2

24 and 222 13 and 73 NR Greve (1985).

Iowa 1

25 48 NR Hill and others (1991).

Kansas NR NR NR Lindquist (1978).

Northeast Kansas 128 44 (1–263) Robel and others (1989).

Fort Riley 36 33 14

(1–124)

Robel and others (1989).

Rural Manhattan 92 75 21

(1–263)

Robel and others (1989).

Eastern Kansas 8 5 of 8 NR Ball and others (1998).

Michigan 256 0 NR Stuewer (1943).
Causative Agent 17

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

Midwestern United States—Continued

Michigan, Wisconsin, and Ohio 25 32 (1–18) R.L. Rausch, unpub. data 

(1944–48).

Southeast Michigan 33 58 27

(1–110)

Schultz (1962).

Michigan 1 1 of 1 NR Thomas (1988).

Minnesota 1 1 of 1 Numerous Olsen and Fenstermacher (1938).

Minnesota 1 0 NR Larson and Scharf (1975).

Minnesota 163 61 NR S. Schmidt and others,

unpub. data (1998).

Twin Cities Metro Area 109 66 NR S. Schmidt and others,

unpub. data (1998).

Nebraska 4 3 of 4 NR Armstrong and others (1989).

Southern Ohio 1 1 of 1 NR Rausch (1946).

Central Ohio 28 25 NR Dubey (1982).

Southwest Ohio 226 54 14

(NR–210)

Ingle and others (2014).

South Dakota 250 12 NR

(NR–46)

Boddicker and Progulske (1968).

West Virginia (far north) 10 20 3

(NR-5)

Schaffer and others (1981).

West Virginia (central) 3 0 NR Owen and others (2004).

West Virginia (central) 1

22 0 NR Owen and others (2004).

Wisconsin 213 51 NR

(1–241)

Amundson and Marquenski 

(1986).

 Southern half 114 75 48 Amundson and Marquenski 

(1986).

 Northern half 99 23 28 Amundson and Marquenski 

(1986).

Southern Wisconsin 57 65 NR Samson and others (2012).

Northeast and Middle Atlantic United States

Connecticut 1 1 of 1 (1,321) Carlson and Nielsen (1984).

Maryland 19 5 Many Habermann and others (1958).

Maryland 1

304 30 NR K.R. Kazacos and N. Garner,

unpub. data (1990).

New Jersey 1

21 24 NR LoGiudice (1995).

New Jersey 2

137 34 NR LoGiudice (1995).

New Jersey 54 67 15

(1–58)

LoGiudice (2001).

New Jersey 2

208 22 NR LoGiudice (2003).

New York City 1 1 of 1 (10) McClure (1933).

New York City NR Positive NR Herman (1939).
18 Baylisascaris Larva Migrans

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

Northeast and Middle Atlantic United States—Continued

Eastern New York 2 2 of 2 389

(141–636)

Stone (1983).

Western New York 429 68 48

(1–480)

Ermer and Fodge (1986).

Southern New York (Ithaca) 1

277 20

(4–42)

NR Kidder and others (1989).

Southeast New York

(Long Island)

2

49 39 NR Feigley (1992).

Western Pennsylvania 52 58 14

(1–170)

Noce and others (2007).

Southeast Pennsylvania 1 1 of 1 5 Dubey and others (1992).

Pennsylvania

2010 189 38 NR Cottrell and others (2014).

2011 1383 33 NR Cottrell and others (2014).

Washington, D.C. 23 35 NR Tecec (1987).

Washington, D.C. 2

21 52 NR Tecec (1987).

Southeastern United States

Alabama 371 0 NR Johnson (1970).

Arkansas 30 0 NR Richardson and others (1992).

Florida 19 0 NR Harkema and Miller (1964).

Florida 158 0 NR Forrester (1992).

Central Florida 51 0 NR Schaffer and others (1981).

Southeast Florida

(Miami)

90 0 NR K.R. Kazacos and others,

unpub. data (1997).

Southeast Florida

(Key Largo)

1

64 0 NR McCleery and others (2005).

Southeast Florida

(Broward County)

1 1 of 1 Several Blizzard, Yabsley, and others 

(2010).

Northwest Florida

(Leon and Wakulla County)

3 3 of 3 1 Blizzard, Yabsley, and others 

(2010).

Northwest Florida (Bay,

Escambia, Leon County)

52 12 NR D. Wolf, written commun. (2013).

West-central Florida

(Hillsborough, Pasco,

Pinellas County)

183 18 NR D. Wolf, written commun. (2013).

East-central Florida

(Orange County)

25 16 NR D. Wolf, written commun. (2013).

Central Georgia 6 Positive NR Babero and Shepperson (1958).

Eastern Georgia

(Ossabaw Island)

100 0 NR Jordan and Hayes (1959).

Georgia 22 0 NR Harkema and Miller (1964).
Causative Agent 19

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

Southeastern United States—Continued

Georgia 312 4 NR Blizzard, Davis, and others 

(2010).

Northern Georgia

(Clarke County)

116 10 7

(1–22)

Blizzard, Davis, and others 

(2010).

Northern Georgia 110 1 NR V.R. Nettles, unpub. data 

(1976–77).

Northeast Georgia

(Atlanta)

50 22 6

(1–24)

Eberhard and others (2003).

Northern Georgia and western North 

Carolina

23 0 NR Schaffer and others (1981).

Southeast Georgia 10 0 NR Schaffer and others (1981).

Eastern Georgia

(St. Catherine Island)

32 0 NR Price and Harman (1983).

Western Kentucky 70 30 NR

(NR–61)

Cole and Shoop (1987).

North Carolina

 Coastal 61 0 NR Harkema and Miller (1964).

 Inland 148 0 NR Harkema and Miller (1964).

Southeast North Carolina 10 0 NR Schaffer and others (1981).

Western North Carolina 74 12 20

(1–122)

Hernandez and others (2013).

South Carolina (Cape Island) 16 0 NR Harkema and Miller (1962).

South Carolina

 Cape Island 17 0 NR Harkema and Miller (1964).

 Coastal 16 0 NR Harkema and Miller (1964).

 Inland 31 0 NR Harkema and Miller (1964).

Tennessee 253 8 (1–221) Bafundo and others (1980).

Northeast Tennessee/Virginia 20 0 NR Schaffer and others (1981).

Eastern Tennessee 118 13 7

(1–15)

Souza and others (2009).

Northwest Tennessee and

southwest Kentucky

145 3 NR

(NR–83)

Smith and others (1985).

Virginia 6 0 NR Harkema and Miller (1964).

Western Virginia 7 5 of 7 (17–93) Jacobson and others (1976).

Eastern Virginia (coastal) 10 0 NR Schaffer and others (1981).

Eastern Virginia 38 0 NR Jones and McGinnes, (1983).

Western Virginia (mountains) 34 56 16

(NR-129)

Jones and McGinnes (1983).

Northern Virginia 44 2 NR Hancock and others (2002).
20 Baylisascaris Larva Migrans

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

West and Southwest United States

California NR Positive NR Voge (1956).

Southern California NR Positive NR Overstreet (1970).

Southern California (Santa Barbara) 26 92 NR Moore and others (2004).

Northern California 12 67 NR Goldberg and others (1993).

Northern California 26 58 NR Park and others (1998).

Northern California 56 70

(52 and 1

18)

(1–tremendous

numbers) 

W.J. Murray and others,

unpub. data (1999).

Northern California (coastal) 15 100 NR W.J. Murray and others, unpub. 

data (1998).

Northern California (coastal) 2

215 44–53 NR Rouserre and others (2003).

Northern California (Marin,

Sonoma, Alameda County)

10 60 3

(1–10)

Kimball and others (2003).

Northern California (Marin,

Sonoma, Alameda County)

2

39 67 NR Kimball and others (2003).

Eastern Colorado 53

(46 and 1

7)

59

(57 and 171)

12

(1–49)

Chavez and others (2012).

Colorado 1

1 1 of 1 NR Markanzky (2015).

Western Nevada (Reno) 4 3 of 4 NR Murray and others (2005).

Western Nevada (Reno) 2

38 37 NR Murray and others (2005).

Oklahoma 1 1 of 1 (Large

numbers)

Campbell and others (1997).

Oregon 1 0 NR Senger and Neiland (1955).

Northwest Oregon (Portland) 69 58 NR Yeitz and others (2009).

Eastern Texas 13 0 NR Chandler (1942).

Central Texas 37 0 NR Schaffer and others (1981).

Eastern Texas 1

62 23 NR S.C. Waring and D.D. Dingley,

written commun. (1989) in

Kazacos and Boyce (1989).

Southern Texas (coastal,

Corpus Christi)

33 70 6

(1–28)

Kerr and others (1997).

Southern Texas 19 16 3

(1–4)

Long and others (2006).

Southern Texas (Kingsville) 55 5 7

(6–9)

Reed and others (2012).

Texas 590 5 7 Kresta and others (2009, 2010). 

Utah 1

152 35 NR B. Zscheile and others, unpub. 

data (2011).

Utah 35 NR 12

(1–57)

B. Zscheile and others, unpub. 

data (2011).

Southwest Washington 29 3 1 McNeil and Krogsdale (1953).

Washington 62 79 NR W.J. Foreyt, written commun. 

(1998).
Causative Agent 21

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

West and Southwest United States—Continued

Western Washington 1

5 1 of 5 NR Markanzky (2015).

Wyoming 363 45 11

(1–88)

Pipas and others (2014).

Canada

Southwest British Columbia 82 61 27

(1–226)

Ching and others (2000).

Southeast Manitoba (Winnipeg) 114 50 NR Sexsmith and others (2009).

New Brunswick 117 21 (1–97) S. Scott and others, unpub. data 

(2010); M. Duffy, written

commun. (2010).

Nova Scotia 219 8 NR Anderson and Mills (1991).

Nova Scotia 236 7 (1–46) Smith (1992).

Nova Scotia 491 7 NR J. Mills, unpub. data (1993).

Ontario NR Positive NR Sprent (1968).

Ontario (Toronto) 23 43 Numerous Cranfield and others (1984).

Southern Ontario (Guelph) 41 51 (1–40) Berry (1985).

Ontario 128 38 3

(1–116)

Jardine and others (2014).

Prince Edward Island 50 2 NR G.A. Conboy, unpub. data (1998).

Prince Edward Island 242 7 NR Conboy and others (2010).

Quebec 21 57 (34–86) Mackay and others (1995).

Southern Saskatchewan 31 0 NR Hoberg and McGee (1982).

Europe

Czech and Slovak Republics 1 1 of 1 2 Tenora and others (1991).

Czech and Slovak Republics 1 1 of 1 28 Tenora and Stanek (1990).

Denmark 1

19 11–21 NR Brinch (2006).

Germany

 Brandenburg 41 0 NR Lux and Priemer (1995).

 Hessen 185 71 (1–232) Gey (1998).

 Bad Karlshafen 1

15 80 NR Hoffmann and others (2002).

 Saxony 56 39 NR Winter and others (2005).

Netherlands NR NR Present H. van Bolhuis, written commun. 

(2009). 

Norway 4 4 of 4 53

(11–115)

Davidson and others (2013).

Poland 1 1 of 1 50 Stefański and Žarnowski (1951).

Poland 2

27 4 NR Bartoszewicz and others (2008).

 Warta Mouth National Park 1

91 3 NR Popiotek and others (2011).

Spain 3 0 NR Jimenez Martinez and others 

(2015).
22 Baylisascaris Larva Migrans

Table 5. Location, prevalence, and numbers of intestinal Baylisascaris procyonis in raccoons.—Continued

[Each line in the table represents a separate study or examination; <, less than; ~, approximately; NR, not reported; Positive, percent unknown]

Location

Number of

animals

examined

Percent

(or number

if <10)

of animals

infected

Mean number

and range of

parasites found

in host animal(s)

(range in

parentheses)

Reference

Asia

Japan 1

291 27 NR Miyashita (1993).

 21 zoos 1

178 40 NR Miyashita (1993).

 6 animal dealers 1

37 8 NR Miyashita (1993).

 Pets 1

39 8 NR Miyashita (1993).

 Wild 1

37 0 NR Miyashita (1993).

 Wild 140 0 NR Kawanaka and others (2001).

 48 zoos 1

48 zoos 13 NR Kawanaka and others (2002).

 Wildlife park 12 33 56

(2–149)

Sato and others (2003).

 Wildlife park 1

12 25 NR Sato and others (2003).

 Wild 531 0 NR Sato and Suzuki (2006).

 Wild 1,688 0 NR Matoba and others (2006).

China

 18 zoos 1

277 13 NR Xie and others (2014).

1

 Examination of raccoon feces for eggs.

2

 Examination of raccoon feces from latrines for eggs.
No data

Reported in isolated areas 

or no prevalence data 

noted

Reported in low prevalence 

(<25 percent)

Reported in moderate 

prevalence (25−60 

percent) in various areas 

of the state or province

Reported in high prevalence 

(> 60 percent) in some or 

most areas of the state 

or province

Distribution of Baylisascaris 

procyonis

EXPLANATION

Data source: Kazacos 2014

CANADA

MEXICO

Geographic Distribution 23

Figure 5. Distribution and general prevalence of Baylisascaris procyonis in raccoons in the United States and Canada. (Modified from 

Hernandez and others, 2013)

Until recently, the farthest known southeastern distribution of B. procyonis was in central Georgia (Babero and 

Shepperson, 1958), but the parasite was recently found in 

raccoons in Florida (Yabsley, 2010; Blizzard, Yabsley, and others, 2010; D. Wolf, written commun., 2013). In the Southeast 

and Mid-Atlantic states, B. procyonis has been found primarily in mountainous areas, not in coastal regions or on coastal 

islands, except in south coastal Texas (Kerr and others, 1997) 

and Florida (Yabsley, 2010; Blizzard, Yabsley, and others, 

2010; D. Wolf, written commun., 2013). It is doubtful that the 

lack of B. procyonis in the deep southeastern United States is 

related to environmental limitations, because Toxocara spp., 

roundworms of dogs, cats, and foxes, and other ascarids do 

very well there (Kazacos, 2001). It likely has more to do with 

expansion of the parasite into new areas, especially past the 

Appalachian mountains, and its establishment and maintenance in the southeast is likely related to host densities and 

other factors. In areas with diverse ecoregions, such as Texas, 

a combination of average rainfall, temperature, soil type, and 

host abundance appear to influence the relative prevalence of 

B. procyonis. Prevalence was higher in central Texas ecoregions and in areas with clayey soil, which retains water better 

and likely helps ameliorate the effects of high temperatures on 

egg survival compared to other soil textures (Kazacos, 1991; 

Kresta and others, 2010). With the introduction, translocation, 

movement, and population increases of raccoons and other 

definitive hosts, B. procyonis has now become established in 

new areas of the South and Southeast. Local prevalences of 

B. procyonis may vary widely and change over time, so it is 

important not to discount its occurrence in a particular area 

until an adequate number of raccoons and other potential 

definitive hosts have been examined. 

B. procyonis has been found recently in areas formerly unaffected, indicating its introduction and (or) spread. 

Although B. procyonis was not previously found in east or 

central Texas (Chandler, 1942; Schaffer and others, 1981), it 

was identified recently in raccoons in these areas as well as in 

south Texas (Kerr and others, 1997; Kresta and others, 2009, 

2010; Reed and others, 2012) (table 5; fig. 5). Variation in the 

prevalence of B. procyonis over time and its presence in isolated “pockets” is well demonstrated in south Texas, where a 

high prevalence in the Corpus Christi area (70 percent) on the 

Gulf Coast differed considerably from that seen in Kingsville 

(5 percent), about 35 miles further south and inland, and Duval 

County (16 percent), even further inland (Kerr and others, 

1997; Long and others, 2006; Reed and others, 2012). B. procyonis was also found at a very low prevalence (0.9 percent) 

in north Georgia in 1976–77 (Kazacos and Boyce, 1989), but 

recently, 22 percent of raccoons in suburban Atlanta (Eberhard 

and others, 2003) as well as 10 percent of raccoons in nearby 
None reported

<10 animals reported

10−49 animals reported

50−100 animals reported

>100 animals reported

Numbers of animals cases

EXPLANATION

Data source Kazacos, 2014

CANADA

MEXICO

3 1

7

27

10

199

1

17

28

9

5

6 1

6

1

1

2

727

35

16

307

146

113

821

12

14

61

37

4

6

6

24 Baylisascaris Larva Migrans

Clarke County (Blizzard, Davis, and others, 2010) were found 

to be infected. B. procyonis had been previously unknown in 

Florida (Forrester, 1992; Kazacos, 2001; McCleery and others, 

2005; K.R. Kazacos and others, unpub. data, 1997), but it was 

recently found in raccoons in four counties in the Panhandle 

region as well as in southeast, west-central and east-central 

Florida (Yabsley, 2010; Blizzard, Yabsley, and others, 2010; 

D.Wolf, written commun., 2013) (table 5). Whether it was 

introduced into these areas by the translocation of infected 

raccoons or dogs or spread naturally is unknown. Limited data 

are available on the prevalence of B. procyonis in the Rocky 

Mountain or Southwestern states, although it was recently 

found in raccoons in Nevada (Murray and others, 2005), Colorado (Chavez and others, 2012), Utah (B. Zscheile and others, 

unpub. data, 2011), and Wyoming (Pipas and others, 2014).

Introduction to new areas has clearly been shown with B. 

procyonis in raccoons in Europe and Asia, and with B. potosis

in kinkajous. Both animal species have been translocated and 

introduced either for their fur (in the case of raccoons) or as 

part of zoo collections and the exotic pet trade. For example, 

kinkajous were imported from the Cooperative Republic of 

Guyana in South America by a breeding facility in MiamiDade County, Florida (Kazacos and others, 2011). Infected 

kinkajous originating from that breeding facility were first 

identified in a pet store in Tennessee. Infected pet kinkajous 

from Guyana were also identified recently in Japan (Taira and 

others, 2013). It remains unknown if this parasite has entered 

the local paratenic host population near the breeding facility 

in Florida or if it can also infect and be spread by raccoons or 

other hosts. 

The distribution of clinical cases of Baylisascaris larva 

migrans (NLM and OLM–DUSN) follows the distribution 

and prevalence of B. procyonis in infected raccoons in North 

America and elsewhere. This is likely related to the pathogenicity of B. procyonis, that is, its ability to produce disease 

in other species, as well as to the widespread distribution of 

raccoons and their ability to live in close association with 

humans and domestic animals. In North America, both animal 

and human cases of Baylisascaris LM are more numerous 

in the Midwestern and Northeastern regions of the United 

States and contiguous Canada and along the West Coast than 

in other areas, although some cases have occurred elsewhere 

(figs. 6 and 7). Fewer cases have been reported from outside 

North America, and these were primarily in areas where 

infected raccoons had been introduced previously and (or) are 

now well established. This trend may be expected to increase 

as both raccoons and the parasite proliferate in new areas and 

awareness of B. procyonis as a disease agent also rises. One 

must also consider other Baylisascaris species as possible 

causes of larva migrans following exposure to other definitive 

hosts, but at this time all cases but a few (greater than or equal 

to 96 percent) have been linked to B. procyonis from raccoons 

(Kazacos, 2001).

Figure 6. Numbers and locations of animal cases and outbreaks of Baylisascaris larva migrans in the United States and Canada.
Neural larva migrans

Ocular larva migrans

Ocular larva migrans, 

location within state 

unknown/not available

Type of larva migrans

EXPLANATION

Data source Kazacos, 2014

CANADA

MEXICO

Geographic Distribution 25

Raccoons have become well established in major areas 

of Europe (fig. 4) and Asia, following their intentional release 

to the wild or escape from fur farms or zoos. Raccoons were 

introduced for hunting and fur into Germany in 1927 and 1934 

(Müller-Using, 1959; Lutz, 1984, 1996; Stubbe, 1990) and 

into the former Soviet Union beginning in 1936; the Soviets 

released 1,243 raccoons into widely scattered areas from 1936 

to 1958 (Aliev and Sanderson, 1966). These programs were 

very successful; in 1964, the raccoon population in the former 

Soviet Union was estimated at 40,000–45,000, and an additional 4,000–5,000 were in Germany. In the 1960s, populations of raccoons were present for a limited time in nine territories of the former Soviet Union, including western regions, 

areas in central Asia, and far eastern territories northwest of 

Japan (Aliev and Sanderson, 1966), although presently they 

exist only in the Caucasus region. Raccoons have proliferated in Germany, Poland, and several other areas and spread 

elsewhere, especially since the mid-1970s, and have now been 

recorded in the wild throughout most of Europe. Established 

or limited populations have now been documented in Spain, 

France, Belgium and the Netherlands eastward to Belarus, 

Ukraine, and Russia (Winter, 2006; Bartoszewicz, 2011; 

Beltrán-Beck and others, 2012; Garcia and others, 2012). Sporadic detections have also occurred in a number of other countries, including England, Norway, Sweden, Denmark, and Italy 

(Beltrán-Beck and others, 2012). Raccoons are also present 

in southern Russia, Georgia, and Azerbaijan, much of Japan 

Figure 7. Locations of human cases of Baylisascaris larva migrans in the United States and Canada.

(Ochiai and others, 2002; Ikeda and others, 2004; Matoba and 

others, 2006) and in zoological parks in China (Xie and others, 

2014). No data exist showing their establishment in the wild in 

China, although this would be expected at some point.

Following the introduction of wild raccoons and their 

subsequent proliferation, it was estimated that by the 1990s 

more than 100,000 wild raccoons were living in Germany 

(C. Bauer, pers. commun., 1991), with a prevalence of 

B. procyonis infection of 71 percent (Gey, 1998). Other 

surveys have also found the parasite at high prevalence in 

Germany, ranging from 39 percent (Winter and others, 2005) 

to 80 percent (Hoffmann and others, 2002). B. procyonis has 

also been identified in raccoons in Poland (Stefański and 

Žarnowski, 1951; Bartoszewicz and others, 2008; Popiotek 

and others, 2011), the Czech Republic (Tenora and Stanek, 

1990), and recently in Denmark (Brinch, 2006), the Netherlands (H. van Bolhuis, written commun., 2009), and Norway 

(Davidson and others, 2013); it will likely be found elsewhere 

as more raccoons are examined. Typical of what happens, 

four raccoons confiscated from a farm in southern Norway 

had been illegally imported (along with raccoon dogs and 

coatis, also alien species) from an undisclosed location, and 

all of the raccoons were moderately infected with B. procyonis. Two additional raccoons escaped, with one later captured 

and killed and the other still at large (Davidson and others, 

2013). The increase and spread of raccoons in Europe has 

been accompanied by B. procyonis-induced larva migrans in 
26 Baylisascaris Larva Migrans

various species, including humans (Schrott, 1961; Kelly and 

Innes, 1966; Koch and Rapp, 1981; Küchle and others, 1993; 

Jimenez Martinez and others, 2015). The occurrences of such 

cases may increase, and the parasite deserves closer scrutiny 

and investigation as a cause of animal and human disease in 

Europe and Asia. 

Japan encountered major problems with B. procyonis, 

stemming from the importation of over 20,000 raccoons from 

North America as exotic pets for 15 years starting in 1977, 

following a new popular television cartoon series, “Rascal 

Raccoon” (Miyashita, 1993; Beltrán-Beck and others, 2012); 

many of these animals escaped and now inhabit wild areas of 

the country (Ochiai and others, 2002; Ikeda and others, 2004). 

In Japan, B. procyonis was found in raccoons kept by wild animal dealers, wildlife parks, zoos, and pet owners (Miyashita, 

1993; Sato and others, 2003) (table 5). A survey of raccoons in 

48 zoos in Japan found that 6 zoos housed raccoons infected 

with B. procyonis (Kawanaka and others, 2002). Related to 

this, several outbreaks of NLM have been documented in 

animals in Japan, including in a rabbitry and a primate facility 

(Sato and others, 2003, 2005), but to date no human cases 

have been reported. The Japanese acted quickly in the early 

2000s to try to eliminate the parasite by testing and euthanasia or treatment of infected raccoons, and the hope is that 

B. procyonis has not become established in the wild in Japan. 

In studies, 140 wild raccoons were examined from Kanagawa 

and Aichi Prefectures (Kawanaka and others, 2001), 531 raccoons from 12 municipalities in Wakayama Prefecture (Sato 

and Suzuki, 2006), and 1,688 free-ranging raccoons from 

the three main islands (Matoba and others, 2006); none were 

found to be infected with B. procyonis. 

China is facing similar problems with B. procyonis. Over 

320 captive raccoons were present in 18 zoological gardens 

throughout China, and, based on fecal examinations, 35 of 

277 (13 percent) were positive for B. procyonis. A questionnaire survey found that most people interacting with these 

raccoons, including animal keepers and veterinarians, were not 

aware of B. procyonis or larva migrans. To date, no animal or 

human cases of larva migrans due to B. procyonis have been 

reported from China nor have raccoons been found in the 

wild (Xie and others, 2014; Y. Xie, written commun., 2014), 

although both will likely occur. Similar to what was seen in 

China, a questionnaire survey in the United States found that 

many people keeping or interacting with raccoons or nonraccoon procyonids had no prior knowledge of raccoon roundworm, did not know it could be transmitted to other species 

including humans, and did not know that captive animals 

could serve as hosts (Parkanzky, 2015).

The finding of Baylisascaris (originally identified as 

B. procyonis) in a kinkajou from Colombia (Overstreet, 1970) 

extended the geographical range of what is now known as 

B. potosis (Tokiwa and others, 2014) into South America, 

where kinkajous are widely distributed. Baylisascaris was also 

found in pet kinkajous in Indiana and Tennessee, in a breeding 

facility in southeast Florida, and in Japan, where the animals had originated from Guyana, South America (Kazacos, 

Kilbane, and others, 2011; Taira and others, 2013). In a survey 

in the United States, both kinkajous and coatis were found 

to be shedding Baylisascaris eggs (Parkanzky, 2015), and 

there is anecdotal evidence of a patent infection in a coati in 

the Netherlands (H. van Bolhuis, pers. comm. to K. Kazacos, 

2012). The species of Baylisascaris occurring in coatis is 

not known, and it is also unknown if B. procyonis will infect 

other nonraccoon procyonids such as coatis and kinkajous 

when they are in areas enzootic for that parasite. It is possible 

but also unknown if B. procyonis or another species occurs 

in raccoons in Central and South America or the Caribbean, 

or in other nonraccoon procyonids in the Americas. Surveys 

of helminths in crab-eating raccoons and coatis in Brazil 

and in Cozumel raccoons did not find any animals infected 

with Baylisascaris spp. (McFadden and others, 2005; Vieira 

and others, 2008). Two cases of human OLM–DUSN with 

large larvae were recently reported from Brazil (Cialdini and 

others, 1999; Amaro, 2000), and transmission from infected 

raccoons, kinkajous, coatis, or other indigenous animals is a 

likely possibility in these cases. The finding of Baylisascaris

spp. in kinkajous and coatis in the United States and Japan has 

raised the question of possible zoonotic exposure of humans 

through contact with these animals in the wild or via the exotic 

pet trade (Kazacos and others, 2011; Taira and others, 2013). 

However, based on the lack of aggressive somatic migration 

including into the brain and failure to produce clinical disease 

in heavily infected monkeys (Tokiwa and others, 2015), B. 

potosis should be considered of lesser zoonotic importance, 

especially for NLM and (or) OLM–DUSN, than B. procyonis

and some other species. Additional surveys using molecular methods (Tokiwa and others, 2014) are needed to better 

understand the identities and distribution of Baylisascaris

species occurring in various procyonids in the Americas 

and elsewhere.

Patterns and Trends

Baylisascariasis has progressed from a parasitic curiosity to a well-known animal and human disease in a relatively short span of 40–50 years. Awareness of the parasite, 

especially as a zoonosis, has increased dramatically in the 

past 25 years, as more animal and human cases have been 

described and publicized. Both recognition and treatment of 

the first few human cases met with difficulty for three important reasons:

1. medical schools in North America had or have eliminated much of the parasitology training from their 

curricula, based on the premise that parasites were (and 

for many, still are) primarily “foreign” disease problems 

that have been eliminated from North America (Nelson, 

1983); 

2. the average physician received or receives little if any 

training in zoonotic diseases (Kahn, 2006; Kahn and 

others, 2007); and 
Patterns and Trends 27

3. the parasite simply was not known to the medical 

community, even to medical pathologists, so it was not 

considered a possible cause of human disease (Huff and 

others, 1984; Kazacos, Vestre and Kazacos, 1984; 

Sanders, 2015). 

In the first few human cases, it took considerable time 

to rule out a long list of other possible infectious and parasitic causes and to eventually consider Baylisascaris, if at 

all, as being involved. Thus, there were unfortunate delays in 

diagnosis and treatment, even though potentially useful drugs 

were available and being used for toxocaral larva migrans. In 

one case, Baylisascaris was not even considered as a possible 

cause of progressive neurologic disease in an infant until several years later, when characteristic ocular lesions of DUSN 

were noted during a routine eye exam and prompted the diagnosis of Baylisascaris by the use of serologic tests, which analyze the blood serum for antibodies (Gavin and others, 2002; 

Mets and others, 2003). Unfortunately, due to the amount of 

damage to the child’s brain, treatment at that time was without 

benefit and the patient remained profoundly impaired until his 

death from complications years later. Several early patients 

were subjected to open brain biopsies in an attempt to reach 

a diagnosis, and larval sections were fortuitously identified in 

brain tissue (Cunningham and others, 1994; Rowley and others, 2000; Kazacos, Gavin, and others, 2002). With the present 

availability of serologic tests for Baylisascaris, it is no longer 

necessary for these invasive procedures to be used.

The overall situation for Baylisascaris LM diagnosis 

has improved dramatically in the past several decades, based 

primarily on relentless efforts to educate physicians, veterinarians, and public health personnel about the parasite. 

Baylisascaris is now included in various medical textbooks 

and other infectious disease resources, including Web sites 

from the Centers for Disease Control and Prevention (CDC, 

www.cdc.gov), the Center for Food Security and Public Health 

(www.cfsph.iastate.edu), and the Companion Animal Parasite Council (www.capcvet.org). The CDC launched updated 

Web coverage of Baylisascaris in spring 2011 and, along with 

the National Reference Centre for Parasitology in Montreal, 

Canada, took over serologic testing for the parasite, based on 

a recently developed recombinant antigen (Dangoudoubiyam and others, 2010; Rascoe and others, 2013). The current 

situation is still one of inadequate awareness of Baylisascaris

by front-line physicians such as pediatricians and general 

practitioners, but good awareness on the part of infectious 

disease specialists, who are often called in to consult on difficult cases; ophthalmologists, who are well aware of OLM 

and DUSN; and veterinarians, who deal with animal cases and 

have considerable training in zoonotic diseases. Today, if a 

child in North America is diagnosed with eosinophilic meningoencephalitis and an infectious disease expert is consulted, 

Baylisascaris will be very high on the differential list, and 

not only will serum and (or) CSF samples be sent for testing 

within days, but often the patient will be started on immediate 

aggressive anthelmintic and steroid treatment (K.R. Kazacos, 

pers. observation, 2008–15; Hajek and others, 2009; Haider 

and others, 2012; Peters and others, 2012; Sanders, 2015). 

Diagnosis and treatment of human cases will continue to 

improve as additional cases are documented and the medical 

profession gains greater awareness of the infection. 

The risk of human and animal infection with Baylisascaris will continue to increase as populations of raccoons 

expand in close proximity to humans and become established 

in new areas through dispersal and translocation. Because they 

are extremely adaptable, raccoons do well in peridomestic

urban and suburban environments, which record some of the 

highest population densities of these animals (Hoffman and 

Gottschang, 1977; Kidder, 1990; Rosatte and others, 1991; 

Feigley, 1992; Gehrt, 2003; Prange and others, 2003; Roussere and others, 2003; Page, 2013). In these environments, 

raccoons have few natural enemies and benefit from excellent food availability, shelter, and den sites, allowing them 

to flourish in close association with humans. They have truly 

become an urban and suburban wildlife species, and most 

cities and towns in the animal’s range support large numbers 

of raccoons. Urban raccoon populations carrying B. procyonis

have been documented recently in Chicago, Atlanta, the Twin 

Cities metroplex in Minnesota; Portland, Oregon; Orange 

County, California; and Toronto (Kazacos, 2001; Evans, 

2001; Eberhard and others, 2003; Page, Gehrt, and others, 

2009; Yeitz and others, 2009). Unfortunately, along with these 

animals come their feces, to which both humans and animals 

are routinely exposed in urban and suburban environments. 

This, combined with the high prevalence of B. procyonis in 

raccoon populations, makes the likelihood of human exposure 

and infection with this parasite high (Kazacos, 2001; Page, 

Anchor, and others, 2009). 

Many cases of Baylisascaris encephalitis in humans and 

animals are seen in suburban and even urban areas of major 

cities, where it is not unusual to find raccoon latrines around 

peoples’ homes, in their yards, in children’s play areas, parks, 

and other sites with significant human contact (Kazacos, 

2001; Roussere and others, 2003; Page, Anchor, and others, 

2009; Page, 2013). Indeed, three cases of B. procyonis larva 

migrans, one involving an infant with severe NLM and two 

being teenagers with OLM–DUSN, were documented recently 

from Brooklyn and the Bronx, in the urban heart of New York 

City, where raccoons are commonly seen (Perlman and others, 

2010; Saffra and others, 2010; Liu and others, 2015). The 

recent finding that B. procyonis will also mature in some other 

animal species, including domestic dogs, further increases 

possible human exposure from lesser known animal sources 

(Greve and Obrien, 1989; Kazacos, 2006; Kazacos, Kilbane, 

and others, 2011) (table 6). Because of the seriousness of this 

infection and the limitations of diagnosis and treatment, prevention of infection with Baylisascaris is of obvious importance. In addition, greater efforts may be useful to prevent the 

spread of the parasite to new areas, both within and outside 

North America.
28 Baylisascaris Larva Migrans

Table 6. Location, type of infection, number of animals infected, and numbers of intestinal Baylisascaris procyonis, B. potosis, or 

Baylisascaris spp. in nonraccoon definitive hosts.

[NR, not reported] 

Location Type of 

infection

Number of 

animals 

infected

Mean number and 

range of parasites 

found in host animal(s) 

(range in parentheses)

Reference

Kinkajou1

Colombia Natural NR 13 Overstreet (1970).

Indiana Natural 2 (20 to more than 25) Kazacos and others (2011).

Tennessee 2

Natural 1 NR Kazacos and others (2011).

Miami-Dade County, 

Florida

3

Natural NR NR Kazacos and others (2011).

California 2

Natural 2 NR Parkanzky (2015).

Tokyo Natural 1 NR Taira and others (2013).

Northern (bushy-tailed) olingo

NR NR 1 1 Overstreet (1970).

White-nosed coati

Arizona 2

Natural 2 NR Parkanzky (2015).

Netherlands 2

Natural NR NR H. van Bolhuis, 

written commun. (2012).

Domestic dog

Iowa Natural 2 (2–3) Greve and O’Brien (1989).

Missouri Natural 2 13 (NR) Averbeck and others (1995).

Indiana Natural 13 4

7 (2–13) K.R. Kazacos, unpub. data (1999).

Michigan Natural 5

7 and 5 62 (1–3) D.D. Bowman, unpub. data (1998).

Natural More than 17 

(includes above)

NR Bowman and others (2005).

Natural 5 15 (1–40) Bowman and others (2005).

Experimental 7

2 2 (1–2) Bowman and others (2005).

Experimental 7

8 5 (1–14) Reinemeyer and others (2008).

Prince Edward Island, 

Canada

Natural 5 NR (NR–17) Conboy and others (2010).

Natural 2

2 of 555 NR Conboy and others (2010).

Japan Experimental 8

1 of 3 6 Miyashita (1993).

Experimental 7

3 of 4 (4–5) Miyashita (1993).

Virginia opossum

Georgia Experimental 8

0 of 1 NR V.R. Nettles, unpub. data (1976–77), 

written commun. (1985), 

in Kazacos and Boyce (1989).

Experimental 7

1 of 1 13

1

 Parasite redescribed as new species, B. potosis (Tokiwa and others, 2014).

2

 Based on identification of eggs in fecal samples.

3

 Based on identification of eggs in soil samples under cages.

4

 Based on 10 dogs.

5

 Probable infections; based on single fecal exam.

6

 Based on four dogs.

7

 Fed third-stage larvae.

8

 Fed infective eggs.
Patterns and Trends 29

Other Procyonids

As a group, members of the taxonomic family Procyonidae (procyonids) are relatively small, omnivorous

mammals, including raccoons, coatis, ringtails, cacomistles, kinkajous, and olingos (A). Like the raccoon, most 

other procyonids have distinct masklike facial markings 

and banded tails. The primary definitive host of B. procyonis is the common raccoon. It is possible that other raccoon species, such as the crab-eating raccoon and Cozumel 

raccoon, or related procyonids could become infected with 

B. procyonis, although this is not well known. The extent 

of cross infectivity of B. procyonis for other procyonids 

and the exact species involved in some cases of infection 

remain to be determined by molecular and other methods.

Patent intestinal infections with Baylisascaris sp. 

have been documented multiple times in kinkajous, one 

captured in Colombia, South America (Overstreet, 1970) 

and others kept as pets or in breeding facilities, zoos, or 

animal rescue/wildlife sanctuary facilities in the United 

States (Yabsley, 2010; Kazacos, Kilbane, and others, 2011; 

Parkanzky, 2015) and Japan (Taira and others, 2013). 

These infections were thought to involve B. procyonis, but 

the parasite in kinkajous was recently described as a new 

species, B. potosis (Tokiwa and others, 2014). Kinkajous 

are routinely imported from South America and (or) bred 

for sale as exotic pets in the United States and elsewhere. 

B. procyonis infection was also documented in a Northern olingo, but researchers were unsure if it resulted from 

natural or experimental infection (Overstreet, 1970) and 

the actual identity of the parasite is open to question. Eggs 

of B. procyonis or a related species were also found in the 

feces of coatis in the United States (Parkanzky, 2015) and 

the Netherlands (H. van Bolhuis, written commun., 2012). 

Coatis, ringtails and the other procyonids live in the wild 

in the Americas and may be imported, raised, and sold as 

exotic pets, similar to kinkajous. More attention could beneficially focus on all of these animals as additional potential 

sources of Baylisascaris infection for animals and humans 

in North and South America or wherever else they may be 

imported, raised, or sold. If the animals are kept as pets, 

owners and family members will have considerable direct 

contact with them as well as their potentially contaminated 

cages or enclosures, similar to keeping pet raccoons. The 

animal sources in cases of human OLM–DUSN caused by 

a large nematode and recently reported from South America 

(Cialdini and others, 1999; Amaro, 2000) are presently 

Other Hosts of Adult Baylisascaris procyonis

A. Procyonid hosts of Baylisascaris spp.

1. Crab-eating raccoon (Photo by Steven 

G. Johnson), 2. Cozumel raccoon (Photo 

by Camazine), 3. White-nosed coati 

(Photo by Jongleur 100), 4. Ringtail 

(Photo by Robertbody at en.wikipedia), 

5. Cacomistle (Photo by Autosafari), 

6. Kinkajou (Photo by Robrrb at 

en.wikipekia), 7. Northern or bushy-tailed 

olingo (Photo by Jeremy Gatten)

1

2

3

4

5

6

7
30 Baylisascaris Larva Migrans

Other Hosts of Adult Baylisascaris procyonis

unknown and may involve contact with one or more of 

these animal species infected with B. procyonis or related 

Baylisascaris species.

Domestic Dog

Adult B. procyonis parasites have also been found in 

the domestic dog, which is a troubling association given the 

dog’s importance as a working, service, and companion 

animal. Patent infections with B. procyonis were first seen 

in dogs in Iowa in 1988 (Greve and O’Brien, 1989), and 

since then dozens of cases have been identified in domestic 

dogs from the Midwest and other parts of the United States 

and Canada where raccoons are common (Kazacos, 2001, 

2006). In several of these cases, B. procyonis occurred as a 

mixed infection with Toxocara canis and other helminths

(B, C).

It is not known how these particular dogs became 

infected with B. procyonis; however, in experimental studies, one of three dogs fed infective eggs and three of four 

dogs fed third-stage larvae from mice developed intestinal 

B. procyonis infections (Miyashita, 1993). Patent infections 

were also produced in dogs fed eggs or larvae as part of 

trials of deworming drugs in the United States (Bowman 

and others, 2005; Reinemeyer and others, 2008). It appears 

from experimental studies that tissue larvae are more 

efficient than eggs for inducing patent intestinal infection 

in dogs. In addition, based on the very young age of several 

infected dogs, there is circumstantial evidence of prenatal 

(transplacental) infection in dogs, similar to the situation 

with Toxocara canis (Kazacos, 2006). There is already a 

precedent for prenatal infection with B. procyonis larvae—

larva migrans was documented in newborn lambs in Idaho 

(Anderson, 1999), thus indicating the role of hormonal 

activation of larval migration with Baylisascaris that is well 

known with Toxocara in dogs. 

The biological relationship of B. procyonis and dogs is 

particularly interesting when one considers that, in addition to supporting patent intestinal infections, several dogs 

have died from severe neural larva migrans (NLM) due to 

this parasite (Snyder, 1983; Thomas, 1988; Rudmann and 

others, 1996; H.R. Galano and others, unpub. data, 2005). 

Whether dogs with patent infections also have larvae in 

their somatic tissues, or vice versa, is not known, because 

NLM and patent infections have not been documented in 

the same dog. Differences in how the parasite behaves in 

dogs may be related to how dogs become infected, as well 

as their age, immune status, and prior infection history 

at the time of infection, including prior infection with T. 

canis, the dog roundworm.

Dogs infected with B. procyonis pose a definite 

zoonotic threat because they can contaminate the domestic 

environment with eggs and would not be suspected as a 

source of B. procyonis. Because dogs are indiscriminate 

defecators, contamination from them would be more widespread than from raccoons, which favor focal latrine sites 

(Kazacos, 2001, 2006). Because pet dogs often travel with 

their owners, infected dogs could also spread the parasite to 

new geographical areas. If pet dogs can support B. procyonis, then another open question is whether the parasite can 

become patent in other canine species, including coyotes 

and wolves. These canids would be at an increased risk of 

contracting the parasite, based on their predatory habits and 

regular ingestion of infected paratenic hosts. At present, 

data addressing this question are limited. Among numerous (n=311) coyotes examined from the Midwest, none 

were found to be infected with B. procyonis (K.R. Kazacos, 

unpub. data, 2005); the primary ascarid found in them was 

Toxascaris leonina, and it is unknown if prior infection 

with that parasite would somehow prevent infection with 

B. procyonis.

B. A young dog found to be 

infected with both B. procyonis

and Toxocara canis. Animals 

with intestinal infection usually 

do not show clinical signs. 

(Photo by Sam Royer) 

C. Eggs of Toxocara canis (left) 

and B. procyonis (right) from 

the dog with a dual infection. 

(Photo by Kevin Kazacos)
Species Susceptibility 31

Opossum

Another animal that needs greater examination is the 

Virginia opossum (D). An attempt was made to infect two 

opossums using either infective eggs or tissue larvae of 

B. procyonis (V.R. Nettles, unpub. data, 1976–77, written commun., 1985, cited in Boyce and Kazacos, 1989). 

Although the egg inoculation failed, an intestinal infection 

was established in the opossum fed tissue larvae. There 

have been anecdotal reports of Baylisascaris eggs present 

in the feces of opossums at wildlife rehabilitation centers; 

however, these cases were not followed up with deworming 

to isolate and identify any parasites present. Consequently, 

it is unknown if these reports represent eggs merely passing 

through the intestinal tract resulting from opossums feeding 

on feces at raccoon latrines (Page and others, 1999, 2001a; 

Smyser, Page, and others, 2013) or true intestinal infection D. Virginia opossum. (Photo by U.S. Fish and Wildlife Service)

with B. procyonis. It has become apparent that development 

of B. procyonis in nonraccoon hosts is a definite possibility 

and must be examined in greater detail. 

Other Hosts of Adult Baylisascaris procyonis

The export and translocation of raccoons has already 

spread B. procyonis to Europe and Asia, where it has become 

an established problem, and prevalence rates in some areas 

are at or approaching those in parts of North America (C. 

Bauer, pers. commun., 1991; Miyashita, 1993; Gey, 1998) 

(table 5). Similarly, importation of infected kinkajous with B. 

potosis from South America, as well as the spread of infected 

raccoons, has introduced both parasites into Florida, which 

had previously been free of Baylisascaris (Kazacos, 2001; 

Blizzard, Yabsley, and others, 2010; Yabsley, 2010; Kazacos, 

Kilbane, and others, 2011; D. Wolf, written commun., 2013). 

Humans and animals in those areas may suffer consequences 

from this exposure, especially from B. procyonis, with cases 

of NLM and OLM similar to those seen in other parts of North 

America; this has already occurred in parts of Europe and in 

Japan (Schrott, 1961; Koch and Rapp, 1981; Kuchle and others, 1993; Sato and others, 2003, 2005; Jimenez Martinez and 

others, 2015).

These are all excellent examples of how better education, regulations, and oversight may be useful concerning the 

movement and prophylactic treatment of wildlife and exotic 

animals to prevent the spread of parasites and infectious 

diseases to new regions of the globe. Frequently, the emphasis 

is on domestic animals and communicable diseases, and not 

on wildlife, exotic animals, or parasites (Kahn, 2006; Centers for Disease Control and Prevention, 2015; U.S. Department of Agriculture-Animal and Plant Health Inspection 

Service, 2015).

Species Susceptibility

Few other parasites are as indiscriminate, or noteworthy, 

as B. procyonis in causing neurologic disease in wild, zoo, 

and domestic animals as well as humans (Kazacos, 2001). 

The parasite has been implicated in causing NLM in over 150 

species of birds and mammals, including humans, in North 

America and elsewhere (tables 1 and 2). B. procyonis is also a 

well-known cause of ocular disease (OLM, DUSN) in humans 

and experimental animals. Clinical larva migrans may also 

occur with some other Baylisascaris species, particularly B. 

columnaris of skunks and B.melis of badgers (table 4).

Human Infections

Baylisascaris infection in humans causes a spectrum of 

disease depending on the dose of infective eggs and the location and extent of tissue damage and inflammation caused 

by migrating larvae. Baylisascaris infection has been found 

by use of serologic tests many times in apparently healthy 

humans (Cunningham and others, 1994; Conraths and others, 1996; Brinkman and others, 2003; C.A. Hall and others, 

unpub. data, oral commun., 2014) and in nonhuman primates 

in zoos (Zimmerman and others, 2010) (table 7), indicating 

that low-level, subclinical, or covert infection is common. This 

would be expected from a ubiquitous organism with a resistant 
32 Baylisascaris Larva Migrans

Table 7. Primates seropositive for Baylisascaris larva migrans.1 —Continued

[>, greater than]

Host Location Number of animals 

affected Reference2

Black-and-white ruffed lemur Missouri 1 Zimmerman and others (2010).

Red-handed (Midas) tamarin Indiana 3

1 M.C. Spriggs, unpub. data (2009).

Mantled guereza 

(Abyssinian black-and-white 

Colobus monkey)

Tennessee 9 Zimmerman and others (2010).

De Brazza’s monkey Indiana 2 Zimmerman and others (2010).

Diana monkey Oklahoma 6 Zimmerman and others (2010).

Francois’ langur Tennessee 1 Zimmerman and others (2010).

Rhesus macaque Wisconsin 8 K.R. Kazacos, unpub. data (2006).

Lion-tailed macaque Tennessee 9 Zimmerman and others (2010).

Hamadryas baboon Tennessee 2 Zimmerman and others (2010).

Olive baboon Georgia 3

1 M.L. Eberhard and others, unpub. data (2002).

White-handed gibbon Tennessee 1 Zimmerman and others (2010).

Northern white-cheeked gibbon Tennessee 4 Zimmerman and others (2010).

Siamang Oklahoma 2 Zimmerman and others (2010).

Tennessee 2 Zimmerman and others (2010).

Western lowland gorilla Tennessee 3 Zimmerman and others (2010).

Sumatran orangutan Tennessee 2 Zimmerman and others (2010).

Bornean orangutan Tennessee 3

1 D. Zimmerman, unpub. data (2005).

Bornean orangutan hybrid Wisconsin 3

1 Hanley and others (2006).

transmission stage and with varying exposure levels affecting 

great numbers of people. Subclinical and covert infections 

would be characterized by low levels of serum antibodies, 

with or without mild eosinophilia, and without signs of VLM, 

OLM, or NLM; such cases could be missed by clinicians 

(Kazacos, 1991, 1997, 2000, 2001; Cunningham and others, 

1994; Conraths and others, 1996). These cases would, however, account for the fact that a background level of Baylisascaris seropositivity would occur in any exposed population, 

similar to the situation for human infections with dog and cat 

roundworms (Won and others, 2008; Lee and others, 2010; 

Fan and others, 2015). 

Baylisascaris infection is most commonly associated with 

encephalitis due to neural larva migrans (Kazacos, 1997, 2000, 

2001). Twenty-five cases of Baylisascaris encephalitis in 

humans have been published or reported (table 8), and many 

additional cases also are known (K.R. Kazacos, unpub. data, 

1995–2010). Baylisascaris encephalitis is seen most often in 

children less than 2 years old (table 8), who are at the greatest risk of infection because of poorer hygiene and greater 

oral sampling of their environment; however, this does not 

preclude the disease in older children or adults if they contract 

a heavy infection. Young children may ingest very large numbers of eggs from raccoon feces or soil at raccoon latrines and 

have sufficient numbers of larvae entering the brain to produce 

severe damage and clinical signs. Because raccoon feces may 

contain tens to hundreds of thousands of eggs per gram, a 

child can ingest surprisingly little material and acquire a heavy 

infection and severe clinical disease from encephalitis. In an 

early fatal case in Illinois, 3 larvae per gram of brain were 

found in an 18-month-old boy during autopsy, resulting in an 

estimated brain burden of 3,207 larvae and an infecting dose 

of 46,000–64,000 eggs (Fox and others, 1985). Such a dose 

of eggs could certainly occur in 1 gram of raccoon feces, and 

a child could easily ingest this amount or more of feces from 

a contaminated area. This child was believed to have become 

infected by sucking or chewing on pieces of contaminated 

firewood bark that contained infective eggs from raccoon feces 

that were present on the original fallen log that was cut up as 

firewood and brought into the home. Most affected children 

have been male, although several recent cases have involved 

infant girls. Other risk factors for infection include developmental delay, mental retardation, dementia, and drug abuse, 

all of which result in altered awareness and behavior and have 

led to heavy infections in adults (Cunningham and others, 

1994; Kazacos, Gavin, and others, 2002; Chun and others, 

2009; Ciarlini and others, 2011; Hung and others, 2012). Such 

individuals often exhibit the same behaviors seen in young 

children at risk of infection, including increased oral sampling 

of their environment and an urge to eat nonfood substances 

(pica) and dirt (geophagia).
Species Susceptibility 33

Table 7. Primates seropositive for Baylisascaris larva migrans.1 —Continued

[>, greater than]

Host Location Number of animals 

affected Reference2

Human California 4

1 Goldberg and others (1993).

3

1 Kazacos and others (2002).

3

1 Rowley and others (2000).

3

1 Park and others (2000).

3

1 Schultz (2002); Murray and Kazacos (2004).

4 K.R. Kazacos and others, unpub. data (2002).

3

1 C. Langelier and others, unpub. data (2015).

Illinois 3

2 Gavin and others (2002); Kazacos and others (2002); 

Mets and others (2003).

3

1 Fox and others (1985).

Louisiana 3

1 Pai and others (2007)

Massachusetts 3

1 Kelly and others (2012); Peters and others (2012).

Michigan 3

1 J.M. Proos and others, unpub. data (1993).

Minnesota 3

2 Moertel and others (2001).

Missouri 3

1 Anderson and others (1975).

2 Anderson and others (1975).

3

1 Mehta and others (2010).

New York 3

1 C.J. Crosley and others, unpub. data (2005); 

Crosley and Kazacos (2005).

3

1 Perlman and others (2010).

3

1 Cunningham and others (1994).

2 Cunningham and others (1994).

4

1 Saffra and others (2010).

4

1 Liu and others (2015).

Oregon 3

1 Chun and others (2009).

3

1 M. Lahr and R.D. Jensen, written commun. (1986), 

in Cunningham and others (1994).

Pennsylvania 3

1 Huff and others (1984).

Georgia 7 C.A. Hall and others, unpub. data, oral commun. 

(2014).

Various states >100 K.R. Kazacos and others, unpub. data (1995–2010).

Germany 4 Conraths and others (1996).

Ontario 3

1 Hajek and others (2009).

3

1 Reilly (2008); Haider and others (2012).

1

 Serology was done at Purdue University by enzyme-linked immunosorbent assay (ELISA) using B. procyonis larval excretory-secretory antigens 

(Dangoudoubiyam and others, 2011), except for Anderson and others (1975 [indirect hemagglutination and bentonite flocculation for Ascaris]), 

Huff and others (1984 [indirect immunofluorescence]), Fox and others (1985 [indirect immunofluorescence and ELISA using embryonated egg antigens]), 

Goldberg and others (1993 [Western blotting]), Conraths and others (1996 [Western blotting]), and C. Langelier and others, unpub. data 

(2015 [Western blotting]).

2

 Cases have been confirmed by the author and information and data are on file with the author.

3

 Clinically affected with Baylisascaris NLM.

4

 Clinically affected with Baylisascaris OLM.
34 Baylisascaris Larva Migrans

Table 8. Cases of Baylisascaris neural larva migrans in humans.

[Partial list; there may be other cases of which the author is not aware. NR, not reported]

Year of onset 

or presentation

Patient 

age

Patient 

sex

Location Risk factors and 

notes Outcome Reference

1973 18 months Female Missouri Frequent geophagia; 

lived on a farm

Weakness, spasticity Anderson and others 

(1975).

1980 10 months Male Pennsylvania Frequent pica; raccoons 

denned in chimneys

Fatal Huff and others 

(1984); Brown and 

Huff (1983).

1984 18 months Male Illinois Pica of contaminated 

firewood bark; 

Down syndrome

Fatal Fox and others 

(1985).

Circa 1985 21 years Male Oregon Geophagia, pica, 

developmental 

disability1

Persistent central nervous 

system deficits, seizures, 

paralysis of one side of 

the body

Cunningham and others (1994).

1990 13 months Male New York Pica, geophagia near 

raccoon latrine on 

a farm; raccoons 

common1

Severe central nervous system 

impairment, wheelchair 

bound, cortical blindness, 

muscular weakness or 

paralysis of one side of the 

body, brain atrophy

Cunningham and 

others (1994).

1993 9 months Male Michigan Soil contact, probable 

pica; raccoons common in area1

Severe central nervous system 

deficits, cortical blindness, 

hearing loss, seizures, brain 

atrophy

J.M. Proos and 

others, unpub. 

data (1993); 

Murray and 

Kazacos (2004).

1993 13 months Male California None noted, but 

geophagia likely; 

raccoons common in 

area and backyard

Severe central nervous system 

impairment, wheelchair 

bound, seizures, blindness, 

weakness or paralysis of one 

side of the body, incontinence, brain atrophy

Rowley and others 

(2000).

1996 6 years 

(onset 

2.5 

years)

Male Illinois Frequent pica and 

geophagia; contact 

with raccoon feces; 

developmental delay

Severe central nervous 

system impairment, wheelchair bound, seizures, incontinence, without speech

Gavin and others 

(2002).

1996 13 months Male Minnesota None noted; exposure 

to pet and feral raccoons1

Fatal Moertel and others 

(2001).

1997 19 months Male Minnesota Developmental delay, 

Klinefelter syndrome

Fatal Moertel and others 

(2001).

1998 11 months Male California Pica and geophagia; 

many raccoons and 

raccoon latrines on 

property

Severe central nervous 

system impairment, seizures, 

profound visual impairment

Park and others 

(2000).

2000 2.5 years Male Illinois Pica and geophagia, 

including from a 

raccoon latrine

Severe central nervous system 

impairment, cortical blindness, coma, marked muscle 

tension

Gavin and others 

(2002); Kazacos 

and others (2002).
Species Susceptibility 35

Table 8. Cases of Baylisascaris neural larva migrans in humans—Continued.

[Partial list; there may be other cases of which the author is not aware. NR, not reported]

Year of onset 

or presentation

Patient 

age

Patient 

sex

Location Risk factors and 

notes Outcome Reference

2000 17 years Male California Marked geophagia, 

including. from a contaminated sandbox; 

severe developmental 

disability

Fatal Kazacos and others 

(2002).

2002 11 months Male California Geophagia, pica from 

a raccoon fecescontaminated area in 

childcare facility1

Severe central nervous system 

deficits, seizures, cortical 

blindness; gradual improvement, but with persistent 

deficits

Schultz (2002); Murray and Kazacos 

(2004); K.R. 

Kazacos, unpub. 

data (2002–4).

2004 4 years Male Louisiana None noted; raccoons 

present in neighborhood1

Mild central nervous system 

deficits, incoordination, 

then full recovery

Pai and others (2007).

2004 15 months Female New York Exposure to soil in front 

yard; raccoons common, latrine on roof1

Severe central nervous system 

impairment, seizures, 

profound cognitive deficits, 

brain atrophy

C.J. Crosley and 

others, unpub. data 

(2005); Crosley and 

Kazacos (2005).

2005 7 years Male Ontario Contact with positive 

raccoon latrines and 

an open sandbox in a 

yard; geophagia; puts 

hands in his mouth; 

autism; attention 

deficit-hyperactivity 

disorder

Improvement with treatment; 

seizures, cortical visual impairment, no speech, brain 

atrophy

Cheney (2005); Chris 

(2005); Hajek and 

others (2009).

2007 17 years Male Oregon Multiple substance 

abuse; neuropsychiatric problems

Gradual improvement with 

significant cognitive deficits 

in memory, processing, 

communication, attention, 

insight

Chun and others 

(2009).

2007 18 months Female Missouri Geophagia, contact with 

contaminated soil in 

a county park where 

raccoons common1

Severe central nervous system 

impairment, seizures, mild 

or partial paralysis, cortical 

blindness, brain atrophy

Mehta and others 

(2010).

2008 12 months Male New York None noted; 

raccoons known in 

neighborhood

Severe central nervous system 

impairment, cortical blindness, lack of cognitive 

function, spastic paralysis 

of corresponding parts on 

both sides of the body, brain 

atrophy

Perlman and others 

(2010).

2008 14 months Male MassachusettsFrequent pica and geophagia; oral contact 

with family members’ 

shoes; raccoons common in area

Improvement following 

treatment; residual partial 

paralysis of the lower limbs, 

language delay, 

brain atrophy

Kelly and others 

(2009, 2012); 

Peters and others 

(2012).
36 Baylisascaris Larva Migrans

Table 8. Cases of Baylisascaris neural larva migrans in humans.—Continued

[Partial list; there may be other cases of which the author is not aware. NR, not reported]

Year of onset 

or presentation

Patient 

age

Patient 

sex

Location Risk factors and 

notes Outcome Reference

2008 14 months Male Ontario Puts hands in his mouth; 

raccoons in the 

immediate vicinity

Severe central nervous system 

deficits, cortical 

blindness, unable to stand

Reilly (2008); 

Haider and 

others (2012).

2009 54 years Male Missouri Mental retardation; eating food scraps from 

public garbage cans1

Fatal Moore (2009); 

Ciarlini and 

others (2011).

 NR 73 years Female British 

Columbia

Alzheimer’s dementia 

that likely predisposed the patient to 

infection

Incidental finding at autopsy Hung and others 

(2012).

2015 63 years Male California Worked under a house 

where raccoons and 

their feces were 

frequently observed. 

Ate lunch without 

washing hands.

Severe progressive central 

nervous system impairment; 

confusion, cognitive deficits, 

incoordination; gradual 

improvement following 

treatment

C. Langelier and 

others, written 

commun. (2015).

1

 K.R. Kazacos, unpub. data (1985–2009); data on file with the author.
Species Susceptibility 37

Clinical Disease in Humans

Human infection with Baylisascaris includes asymptomatic infection, covert infection, visceral infection or 

disease (visceral larva migrans, VLM), central nervous 

system (CNS) infection or disease (neural larva migrans, 

NLM), and (or) ocular infection or disease (ocular larva 

migrans, OLM) (Kazacos, 1991, 1997, 2000, 2001; Sorvillo and others, 2002; Murray and Kazacos, 2004; Gavin 

and others, 2005; Wise and others, 2005; Shafir and others, 

2006; Kazacos and others, 2013; Singaravelu and others, 

2016). The different types vary primarily in the dose of 

infecting eggs and the location and extent of tissue damage and inflammation caused by migrating larvae. The two 

most important clinical problems related to Baylisascaris

infection are CNS disease resulting from brain invasion and 

encephalitis (NLM) and ocular disease caused by invasion 

of the eye (OLM and diffuse unilateral subacute neuroretinitis, DUSN). The more eggs ingested by a person, the greater 

the chances of developing one or both of these conditions, 

due to the increased likelihood of larvae entering the brain 

or eye. If enough eggs are ingested to result in CNS disease, the patient may also show evidence of visceral disease 

(VLM) caused by early larval migration through the liver 

and lungs, and (or) eye disease (OLM) caused by concomitant larval invasion of the eye.

Asymptomatic Infection

Most cases of Baylisascaris infection are the result of 

incidental or low-level infection with eggs and are asymptomatic, that is, they will not result in recognizable clinical 

signs. This is because, in addition to the low levels involved, 

the vast majority (an estimated 93–95 percent) of migrating larvae enter the muscles, connective tissue, or visceral 

organs, where they are walled off in granulomas (Kazacos, 

1986, 1991, 1997, 2001). Once this happens, they settle 

down, become quiescent, and will not usually migrate 

further or cause additional harm. A few larvae may also 

enter the CNS and become encapsulated there, ceasing their 

migration; this is known in animals (Tiner 1953a; Kazacos, 

1997, 2001; Sheppard and Kazacos, 1997) and was recently 

demonstrated in humans (Hung and others, 2012). Humans 

may have incidental CNS invasion without recognizable 

clinical signs more often than is presently known. Such 

individuals may or may not (but often do) develop detectable levels of antibodies that indicate that they are infected, 

but they do not have any or enough larvae in sensitive or 

critical locations such as the brain or eyes to result in serious clinical problems. In essence, these people represent a 

background level of low infection in the population that, 

along with covert infection, may only be identified through 

antibody testing (Cunningham and others, 1994; Brinkman 

and others, 2003; Zimmerman and others, 2010; Rascoe and 

others, 2013).

Covert Infection

Larger numbers of ingested eggs and larvae migrating 

in various tissues may result in nonspecific clinical signs, 

so-called covert or hidden infection. Best described for Toxocara infection (Taylor and others, 1987), covert infection is 

also known to occur with Baylisascaris. With covert infection, more tissue is damaged and inflamed, causing some 

clinical problems that are generally too nonspecific to allow 

a physician to make a correct diagnosis. With Toxocara, 

and probably Baylisascaris, patients’ complaints include 

abdominal pain, cough, headache, sleep disturbances, and 

the like, all of which can occur with or are mimicked by 

other infections or conditions. Depending on the level of 

infection, patients may or may not have an eosinophilia, but 

their blood tests usually will show they are seropositive for 

antibodies (Taylor and others, 1987). Even though they are 

known to occur based on epidemiologic studies, the vast 

majority of these cases will be missed and (or) misdiagnosed 

by clinicians.

Baylisascaris Visceral Larva Migrans

Once someone ingests Baylisascaris eggs, larvae hatch 

in the intestine and begin their migration through the intestinal wall and into internal organs. Damage to the liver and 

lungs can cause migration-induced pathological changes 

and clinical signs including an enlarged liver, fever, cough, 

and difficulty breathing (Kazacos, 1997, 2000; Kazacos 

and others, 1981, 2013; Wyand-Oullette and others, 1983; 

Singaravelu and others, 2016). Patients will often develop 

high numbers of eosinophils in their blood in response to 

larval antigens (Fox and others, 1985; Kazacos, 1997). In 

one case, B. procyonis VLM apparently caused the sudden 

death of a 10-year-old boy in Massachusetts from a nodular 

eosinophilic mass inside the heart (Boschetti and Kasznica, 

1995). Within the mass were the remains of a large (60–70 

micrometer diameter) degenerating ascarid larva.
38 Baylisascaris Larva Migrans

A. Young child with central nervous 

system disease and blindness, infected 

with B. procyonis at a university daycare 

facility in southern California.

B. Young man, severely 

disabled and confined 

to a wheelchair as a 

result of being infected 

with B. procyonis as an 

infant on a dairy farm in 

upstate New York.

C. Young man, severely 

disabled with both 

NLM and OLM–DUSN 

after being infected 

with B. procyonis as a 

child in his yard in the 

Chicago suburbs.

Human cases of Baylisascaris neural larva migrans. (Photos by Kevin Kazacos)

large by comparison and can absorb a certain level of larvae 

and damage before clinical signs become apparent (Hung 

and others, 2012). Clinical signs of Baylisascaris encephalitis can develop as early as 2–4 weeks postinfection and 

progress rapidly in children, who are both smaller in size 

and likely to have ingested a larger number of infective eggs 

than adults (Kazacos, 2000).

Clinical signs in affected humans resemble those seen 

in nonhuman primates suffering from Baylisascaris NLM 

(Kazacos, Wirtz, and others, 1981). There are subtle initial 

changes in behavior or habits, and with increasing neural damage and inflammation, clinical signs may worsen 

quickly (Kazacos, 2000, 2001; Murray and Kazacos, 2004; 

Kazacos and others, 2013). As seen in animals, including 

nonhuman primates, milder disease from lower level infection can manifest as any combination of subtle neurologic 

problems and deficits. Clinical signs in heavily infected 

children are more severe, reflecting extensive damage to 

the nervous system by migrating larvae. Such patients may 

undergo rapidly progressive brain damage that may result in 

coma and death (Kazacos, 2000).

Baylisascaris Neural Larva Migrans and 

Encephalitis

Clinical neurologic disease due to NLM represents 

the high end of the spectrum of Baylisascaris infection, 

in which a person has ingested many eggs and sufficient 

larvae are entering the brain and spinal cord to result in 

clinically significant and potentially life-threatening disease. 

The severity and progression of CNS disease in Baylisascaris encephalitis depends on the number of eggs ingested, 

the number of larvae entering the brain, the location and 

extent of migration damage and inflammation in the brain, 

and, to some extent, the size of the brain (Kazacos, 1996b, 

1997, 2000, 2001; Kazacos and others, 2013; Singaravelu 

and others, 2016). Thus, clinical disease varies from mild 

or insidious, slowly progressive CNS disease with subtle 

clinical signs, to severe, rapidly progressive disease with 

marked clinical signs (Kazacos, 2001; Murray and Kazacos, 

2004). Even though a single larva in the brain of a mouse 

or small bird can be fatal, the brain of a human is quite 

Clinical Disease in Humans
Species Susceptibility 39

Children who survive may have profound neurologic 

impairment, with muscular weakness or paralysis, blindness, seizures, incontinence, and severe developmental 

delays. They may be incapacitated, wheelchair-bound, or in 

a chronic vegetative state (Kazacos, 2000). Even if they can 

be stabilized initially with aggressive treatment, patients 

may progressively worsen over time as the brain undergoes 

postinflammatory wasting with shrinkage and selective loss 

of white matter (Cunningham and others, 1994; Rowley 

and others, 2000). Signs of VLM may also be present in 

patients with clinical NLM, but VLM signs are greatly 

overshadowed by the signs affecting the nervous system. 

Patients with clinical NLM may also develop lesions of 

OLM due to concomitant larval invasion of the eye.

Baylisascaris Ocular Larva Migrans and 

Diffuse Unilateral Subacute Neuroretinitis

Some patients with Baylisascaris encephalitis also 

develop eye disease, because of increased larval numbers, 

dissemination, and eye invasion (Rowley and others, 2000; 

Park and others, 2000; Gavin and others, 2002; Mets and 

others, 2003; Hajek and others, 2009). However, most 

cases of Baylisascaris-induced eye disease are stand-alone 

cases and do not have any other recognizable clinical signs 

(Goldberg and others, 1993; Saffra and others, 2010; Liu 

and others, 2015). This is because a single larva entering 

the eye can cause serious clinical problems, and this occurs 

by chance, based on accidental ingestion of infective eggs. 

On a population level, the more people who are exposed to 

even low-level infection, which is common, the greater the 

chance of ocular invasion and clinical eye disease developing in some individuals. This explains why more cases of 

OLM and DUSN than cases of clinical NLM are seen in 

humans.

Patients with ocular invasion typically experience 

sudden blurred or loss of vision in one eye, and ophthalmologic examination reveals lesions of DUSN, including 

inflammatory and degenerative changes affecting the retina, 

retinal vessels, and optic nerve head (Gass and Braunstein, 

1983; Goldberg and others, 1993; Saffra and others, 2010). 

These changes may be accompanied by migration tracks 

and infiltrates in the retina and (or) choroid, and sometimes 

granuloma formation (Goldberg and others, 1993; Liu and 

others, 2015). If larvae are seen in the eye and are away 

from sensitive structures, they are usually killed using laser 

photocoagulation (Goldberg and others, 1993; Casella and 

others, 1998; Saffra and others, 2010). Larvae seen in the 

eye can be photographed and measured, and their length 

and other morphologic characteristics can be used to identify them as probable Baylisascaris, separating them from 

other possible parasites (Goldberg and others, 1993).

Anthelmintic treatment of DUSN has also been advocated (Gass and others, 1992), but has had mixed results; 

in several cases anthelmintic treatment was apparently 

successful (Cortez and others, 2005; Tarantola and others, 

2011), while in others it was not effective (Casella and others, 1998).

Clinical signs of neural larva migrans in children.

[From Kazacos (2000)]

Lower level infection Heavy infection

Dullness and somnolence Sudden onset of lethargy.

Tremors or unsteadiness Dullness and irritability.

Confusion Neck rigidity and (or) twisting.

Learning disability Loss of fine motor skills.

Developmental regression Decreased head control.

Inability to sit, stand, or walk 

without assistance.

Impaired vision or blindness.

Eye deviation and (or) rapid 

involuntary eye movements.

Speech deterioration.

Problems with eating.

Increase or loss of muscle 

tone.

Extension of the extremities.

Arching of the body.

Stupor.

Coma.

Death.

Clinical Disease in Humans
40 Baylisascaris Larva Migrans

Table 9. Cases of Baylisascaris ocular larva migrans-diffuse unilateral subacute neuroretinitis in humans.—Continued

[Partial list, there are likely other cases of which the author is not aware; NR, not reported; CLM, cutaneous larva migrans; NLM, neural larva migrans; 

DUSN, diffuse unilateral subacute neuroretinitis]

Year of 

onset or 

presentation

Patient 

age

Patient 

sex

Location Worm

seen?

Length of 

worm in 

micrometers

Notes Reference

1949 25 years Male Florida Yes 1,500 — Parsons (1952).

1960 22 years Female Austria Yes 1,700 Infected raccoons 

present in area1

Schrott (1961).

1974 23 years Male Michigan Yes 1,600 Infected raccoons 

common in area

Raymond and others (1978).

1976 13 years Female Kentucky Yes 2,000 Pet raccoon Raymond and others (1978).

NR 13 years Male Nebraska Yes 1,500–2,000 Infected raccoons 

common in area1

Gass and Braunstein (1983).

NR 65 years Female Illinois Yes 1,500–2,000 Infected raccoons 

common in area1

Gass and Braunstein (1983).

NR 25 years Male Illinois Yes 1,500–2,000 Infected raccoons 

common in area1

Gass and Braunstein (1983).

NR NR Female Illinois Yes 1,500–2,000 Infected raccoons 

common in area1

Gass and Braunstein (1983).

1985 42 years Male Wisconsin Yes 1,260 Outdoorsman; contact 

with raccoon den

Williams and others (1988).

NR 27 years Male Pennsylvania Yes 1,600 Infected raccoons 

common in area

Sivalingam and others 

(1991).

1991 48 years Female Germany Yes 1,500 Pet raccoon Kuchle and others (1993).

Circa 1992 29 years Male California Yes 1,727 Infected raccoons 

common in area1

Goldberg and others (1993).

1995 15 years Male Brazil Yes 1,500–2,000 Nonraccoon 

procyonids and 

skunks in area

Cialdini and others (1999).

1996 6 years Male Illinois No NR NLM with ocular 

lesions of DUSN, 

positive serology

Gavin and others (2002); 

Mets and others (2003).

1997 16 years Female Brazil Yes 1,600–2,000 Dog contact; CLM Amaro (2000).

NR 13 months Male California NR NR NLM with ocular 

lesions suggestive 

of DUSN; positive 

brain biopsy and 

serology

Rowley and others (2000).

1998 11 months Male California No NR NLM with ocular 

lesions of DUSN; 

positive serology

Park and others (2000).

Baylisascaris is also well known as a cause of clinical 

OLM, and at least 28 cases have been recognized in North 

America and Europe, including 24 published reports (table 9). 

OLM and DUSN due to B. procyonis are usually seen in older 

children and adults and are clinically similar to toxocaral OLM 

and DUSN, except that the larvae are much larger and ocular 

damage from larval migration may be more severe. Human 

cases have been linked to keeping pet raccoons (Raymond and 

others, 1978; Kuchle and others, 1993) or having contact with 

wild raccoons and contaminated environments.
Species Susceptibility 41

Table 9. Cases of Baylisascaris ocular larva migrans-diffuse unilateral subacute neuroretinitis in humans.—Continued

[Partial list, there are likely other cases of which the author is not aware; NR, not reported; CLM, cutaneous larva migrans; NLM, neural larva migrans; 

DUSN, diffuse unilateral subacute neuroretinitis]

Year of 

onset or 

presentation

Patient 

age

Patient 

sex

Location Worm

seen?

Length of 

worm in 

micrometers

Notes Reference

1999 16 years Male Ohio Yes 1,500 Infected raccoons 

common in area; 

larva recovered1

L.J. Chorich III and others, 

unpub. data (2000).

2000 2.5 years Male Illinois No NR NLM with ocular 

lesions; positive 

serology

Gavin and others (2002); 

Mets and others (2003).

NR 7 years Male Ontario No NR NLM with ocular 

lesions of DUSN; 

positive serology

Hajek and others (2009).

2001 40 years Male Ohio Yes 1,500 Infected raccoons 

common in area; 

positive serology1

S.M. Gordon and others, 

unpub. data (2001).

NR 50 years Male Illinois Yes 1,563 Infected raccoons 

common in area1

G.A. Fishman and others, 

unpub. data (2001).

Circa 2006 37 years Male Ohio Yes NR Positive serology; 

infected raccoons 

common in area

Brasil and others (2006).

NR 15 years Female New York Yes 1,850 Infected raccoons 

in area; raccoons 

seen in immediate 

vicinity

Saffra and others (2010).

2009 45 years Male Iowa Yes 1,270 Turkey hunter; contact 

with soil positive 

for infective B. 

procyonis eggs1

Tarantola and others (2011).

2009 47 years Male California Yes 1,347 Infected raccoons 

common in area1

D.M. Hirota and others, 

unpub. data (2009).

2009 12 years Female New York No NR Infected raccoons occur in area; positive 

ocular serology1

Liu and others (2015).

2010 41 years Female Connecticut Yes 1,400–1,425 Infected raccoons 

common in area1

Goldberg and Bhatnagar 

(2012).

1

 K.R. Kazacos, unpub. data (1985–2010); data on file with the author.
42 Baylisascaris Larva Migrans

Animal Infections

Raccoons

The common raccoon is a member of the Family Procyonidae, which also includes the crab-eating raccoon of Central 

and South America, the critically endangered Cozumel or 

pygmy raccoon on Cozumel Island in Mexico, coatis, the ringtail and cacomistle, the kinkajou, and olingos, all occurring in 

different parts of the Americas. With regard to raccoons, Baylisascaris procyonis has thus far only been found in the common raccoon, and unless otherwise specified, “raccoon” will 

refer to that species. Prevalence of B. procyonis in raccoons 

varies as a function of the examination technique used (necropsy, fecal examination of individuals or latrines), season, 

predominant land use where the animals live, animals’ ages, 

and other factors (Kazacos, 2001; Page and others, 2005). 

Raccoons infected with B. procyonis usually appear normal 

and do not show any clinical signs, although very heavy infections in young raccoons have occasionally caused intestinal 

blockage and death (Stone, 1983; Carlson and Nielsen, 1984). 

Migration of B. procyonis beyond the intestines, including 

somatic migration, does not appear to occur in raccoons.

Due to age-related differences in susceptibility and 

transmission, B. procyonis typically has a higher prevalence 

in juvenile raccoons (91–94 percent) than in adults (37–55 

percent), although this does not hold true in all surveys. In 

areas of high prevalence, the average number of parasites in an 

animal (the intensity) typically ranges from 43 to 52 worms, 

with juvenile raccoons usually having a higher mean intensity (48–62, range 1–480) than adult raccoons (12–22, range 

1–257); males usually have a higher prevalence of infection 

than females, although this may also vary by study (Snyder 

and Fitzgerald, 1985; Ermer and Fodge, 1986; Kazacos, 

2001; Cottrell and others, 2014; Jardine and others, 2014). In 

some areas, such as the southern United States, prevalences 

and mean intensities of infection may be considerably lower 

(Blizzard, Davis, and others, 2010; Blizzard, Yabsley, and 

others, 2010; Kresta and others, 2010; Reed and others, 2012) 

(table 5). The age distribution of B. procyonis in raccoons 

correlates with what is known about the parasite’s life cycle 

(fig. 8): namely, that young raccoons in their first season are 

very susceptible to egg infection and appear to recruit the 

parasite into the raccoon population and have higher worm 

burdens, egg shedding, and prevalence of infection. 

In some areas, primarily northern temperate regions, 

the prevalence of B. procyonis varies with season (Page and 

others, 2005), and the parasite population undergoes a yearly 

cycle with an apparent self-cure in raccoons during the winter 

months (Kazacos, 2001). Studies involving 1,050 raccoons 

from the Midwestern United States corroborated this, with 

strong evidence of a decrease in the worm population from 

January through June and an increase from June to December 

(Page and others, 2016). Seasonal recruitment of the parasite and its increase in prevalence in juvenile raccoons likely 

contributes to the overall increase in prevalence in raccoons 

seen in an area over the summer and fall months (Kazacos, 

2001; Jardine and others, 2014), although this may also be 

affected by other factors (Page, 2013). As raccoons become 

older, age resistance and (or) intestinal immunity combined 

with spontaneous elimination of the parasites may contribute 

to the lower prevalence of B. procyonis in adults. However, 

adults continue to become infected when they ingest paratenic 

hosts carrying larvae through predation or scavenging (Kazacos, 1983b; Kazacos and Boyce, 1989; Kazacos, 2001; Reed 

and others, 2012), and in some cases by ingesting infective 

eggs (the age dichotomy in transmission method is likely not 

100 percent).

Paratenic Hosts

When it comes to larval infection, Baylisascaris procyonis is a remarkably nonspecific parasite. Larvae can infect a 

surprisingly wide variety of paratenic (transport) hosts. In fact, 

this parasite has one of the widest host infection and diseaseproducing capabilities of any worm parasite known, affecting 

numerous free-ranging and captive wildlife, zoo animals, 

domestic animals, and humans with NLM (Kazacos, 2001; 

Kazacos and others, 2013; Singaravelu and others, 2016). 

Both wild and domestic animals act as paratenic hosts of 

B. procyonis and develop NLM (tables 1, 2, 10–21); however, 

susceptibility to Baylisascaris larva migrans varies among animal groups and species (Wirtz, 1982; Sheppard, 1995, 1996; 

Sheppard and Kazacos, 1997; Kazacos, 2001). Animal groups 

particularly susceptible to Baylisascaris NLM include rodents, 

rabbits, primates, and birds, based on the number of cases and 

species affected. Some animal groups and species appear to be 

less or only marginally susceptible, and parasite migration is 

limited to the intestinal wall or viscera; other groups and species appear to be resistant (Kazacos and Boyce, 1989; Kazacos, 2001). Despite this, one should be cautious concerning 

species susceptibility, because a determination of low susceptibility may simply reflect a lack of recognition of cases, and 

new species records are regularly added to the list of animals 

affected with NLM. However, no cases of B. procyonis NLM 

have been documented in opossums, which are commonly 

exposed when they forage at raccoon latrines (Page, 1998; 

Page and others, 1999), nor in cats, which groom considerably and eat rodents possibly contaminated with eggs and 

(or) containing third-stage larvae. No infections were seen in 

cats fed infective eggs or tissue larvae (Miyashita, 1993; K.R. 

Kazacos, unpub. data, 1981). Until recently, no cases had been 

seen in raptors, which are exposed when they prey on other 

animals or consume carcasses, or in adult hoofstock, which 

are exposed when they consume contaminated hay. Because 

limited or no migration was seen in sheep, goats, and swine 

experimentally infected with B. procyonis (Dubey, 1982; Snyder, 1983; Kazacos and Kazacos, 1984), it was believed that 

ungulates were only marginally susceptible to infection and 

would not develop NLM. However, lambs in Idaho were born 
In humans, larvae migrate and become encapsulated 

in tissues and cause VLM, NLM, and OLM

Adults in small 

intestine

Larvae migrate and become encapsulated 

in tissues and cause VLM, NLM, and OLM

Dogs can also serve 

as definitive hosts 

as they can harbor 

adults and shed eggs

Infective eggs 

ingested

Infective eggs 

ingested

External environment

(~2–4 weeks until infective)

Embryonated

egg with infective 

larva

Eggs

Eggs passed 

in feces

Figure 8. Diagram showing the life cycle of Baylisascaris procyonis. (this caption is just for reference. Actual captions are added during layout in InDesign.)

Small mammals 

(rodents, rabbits, etc.) 

and birds

Species Susceptibility 43

Figure 8. The life cycle of Baylisascaris procyonis.

and should now be considered paratenic hosts. Interestingly, 

shrews were found to be resistant to experimental infection 

with B. procyonis eggs and larvae, at dosages much higher 

than those lethal to mice (Sheppard, 1996); the reasons for 

this resistance are unknown, but may include unique or potent 

gastrointestinal enzymes and (or) failure of eggs to hatch or 

larvae to survive (Kazacos, 2001). Unless complete necropsies 

are done, including a thorough examination of the brain in 

all cases of CNS disease, then apparent species limitations to 

Baylisascaris larva migrans should be regarded with caution.

with Baylisascaris encephalitis (Anderson, 1999), indicating 

not only that larvae can migrate through the body tissues of 

sheep, but that they can move through the placenta (transplacental transfer) of pregnant ewes to infect unborn lambs, 

similar to Toxocara canis in dogs. How often this occurs in 

different mammals is unknown, but transplacental infection 

followed by subsequent parasite maturation in puppies is 

believed to occur in dogs (Kazacos, 2001, 2006). Recently, a 

feedlot steer was diagnosed with Baylisascaris encephalitis 

in Alberta, Canada (E. Janzen, pers. commun., 2009), further 

showing that ruminants can be infected in some instances 
44 Baylisascaris Larva Migrans

Table 10. Rodents with naturally occurring Baylisascaris neural larva migrans.1—Continued

[NR, not reported; >, greater than]

Host Location Probable 

parasite species

Number of 

animals affected Reference

House mouse Illinois B. procyonis Several Tiner (1949; 1953a).

White-footed mouse Illinois B. procyonis 2

1 Tiner (1953a,b; 1954).

Indiana B. procyonis 2

1 and 1 Sheppard and Kazacos (1997); K.R. Kazacos, 

unpub. data (1984).

Indiana B. procyonis 10 of 46 Page (1998).

Deer mouse California B. procyonis 4 of 81 Evans (2002).

Brush mouse California B. procyonis 1 R.H. Evans, unpub. data (2001).

Meadow jumping 

mouse

Ontario B. procyonis 1 D.J. Russell, unpub. data (2004).

Allegheny woodrat Indiana B. procyonis 1 K.R. Kazacos and S.A. Johnson, unpub. data (1996).

Indiana B. procyonis 1 Smyser, Johnson, and others (2013).

New York B. procyonis 10 of 10 and 3

1 McGowan (1993).

New Jersey B. procyonis 1 and 3

2 LoGiudice (2003).

Pennsylvania B. procyonis 1 J. Wright and others, unpub. data (1998).

Dusky-footed woodrat California B. procyonis 6 of 45 Evans (2002).

Muskrat New York B. procyonis 4 W.B. Stone, written commun. (1998).

Ontario B. procyonis 1 B.J. McEwen, unpub. data (1988).

Golden hamster Washington B. procyonis 1 M.M. Garner, written commun. (2010).

California pocket mouse California B. procyonis 1 R.H. Evans, unpub. data (2001).

Eastern chipmunk Indiana B. procyonis 1 K.R. Kazacos and S.A. Johnson, unpub. data (1998).

Eastern gray squirrel Indiana B. procyonis 6 K.R. Kazacos, unpub. data (1997; 2004; 2006).

Illinois B. procyonis 3

2 K.R. Kazacos, unpub. data (1996)

Washington B. procyonis 11 of 16 Tseng (1997).

California B. procyonis 2 of 16 Evans (2002).

Ontario B. procyonis 3

1 S.J. Best, unpub. data (1991).

Ontario B. procyonis 1 I.K. Barker, unpub. data (2003).

Eastern fox squirrel Indiana B. procyonis 2 K.R. Kazacos, unpub. data (1983).

California B. procyonis 5 Stringfield and Sedgwick (1997); C.E. Stringfield 

and others, unpub. data (1995–97).

California B. procyonis 4 M.M. Garner, written commun. (2010).

Red-tailed squirrel Maryland B. procyonis 1 Schueler (1973).

American red squirrel Indiana B. procyonis 1 K.R. Kazacos, unpub. data (1998).

Ontario B. procyonis 3

1 I.K. Barker, unpub. data (2002).

Prevost’s squirrel California B. procyonis 1 and 3

2 M.M. Garner, written. commun. (2010).

Douglas squirrel British 

Columbia

B. procyonis 1 Coates and others (1995).

Thirteen-lined 

ground squirrel 

Illinois B. procyonis or 

B. columnaris4

28 Fritz and others (1968).

Illinois B. procyonis 1 Pigage and others (1983).

Illinois B. procyonis 1 J.I. Everitt and S.E. McDonald, unpub. data (1984).
Species Susceptibility 45

Table 10. Rodents with naturally occurring Baylisascaris neural larva migrans.1—Continued

[NR, not reported; >, greater than]

Host Location Probable 

parasite species

Number of 

animals affected Reference

California ground 

squirrel

California B. procyonis 20 of 119 Evans (2002).

California B. procyonis 1 C.E. Stringfield and others, unpub. data (1995).

Black-tailed prairie dog Iowa B. procyonis NR Greve (1985).

New York and 

Wisconsin 

B. procyonis 3 of 52 Dixon and others (1988).

Ohio B. procyonis Several M.T. Barrie, written commun. (2008).

Illinois B. procyonis 1 K.R. Kazacos and P.L. Wolff, unpub. data (1988).

Ontario B. procyonis 3

1 D.A. Smith, unpub. data (2012).

Woodchuck Pennsylvania B. procyonis or 

B. columnaris4

4 of 4 Richter and Kradel (1964).

Connecticut B. procyonis or 

B. columnaris4

NR Swerczek and Helmboldt (1970).

Virginia B. procyonis 3 of 3 Jacobson and others (1976).

New York B. procyonis or 

B. columnaris4

6 Fleming and Caslick (1978).

New York B. procyonis or 

B. columnaris4

5 of 5 Fleming and others (1979).

Indiana B. procyonis 1 and 3 Kazacos and others (1981); K.R. Kazacos, 

unpub. data (1985–1986).

New York B. procyonis or 

B. columnaris4

12 of 12 Roth and others (1982).

Iowa B. procyonis or 

B. columnaris4

NR Greve (1985).

Kentucky B. procyonis or 

B. columnaris4

1 Lyons and others (2001).

West Virginia B. procyonis 1 Owen and others (2004).

Ontario B. procyonis 1 and 3

5 I.K. Barker, unpub. data (1989–2006).

Ontario B. procyonis 1 and 3

2 B.J. McEwen, unpub. data (1988; 1990; 1992).

Ontario B. procyonis 1 and 3

2 D.A. Smith, unpub. data (1992; 2004)

Western pocket gopher California B. procyonis 1 K.R. Kazacos and F.H. Dunker, unpub. data (1996).

Botta’s pocket gopher California B. procyonis 2 of 2 Evans (2002).

California B. procyonis 2 K.R. Kazacos and F.H. Dunker, unpub. data (1997).

Domestic guinea pig Missouri B. procyonis 30 of 50 Van Andel and others (1995).

Nova Scotia B. procyonis 2 Craig and others (1995).

Patagonian mara Illinois B. procyonis 4 K.R. Kazacos and others, unpub. data (1989).

Capybara Illinois B. procyonis 1 K.R. Kazacos and others, unpub. data (1987).

Long-tailed chinchilla Pennsylvania B. procyonis or

B. columnaris4

6 Richter and Kradel (1964).

Ontario B. procyonis 100 Sanford (1989).

Minnesota B. procyonis 17 of 1,400 A.Wuenschmann and K.R. Kazacos, 

unpub. data (2007).
46 Baylisascaris Larva Migrans

Table 10. Rodents with naturally occurring Baylisascaris neural larva migrans.1—Continued

[NR, not reported; >, greater than]

Host Location Probable 

parasite species

Number of 

animals affected Reference

North American 

porcupine

New Jersey B. procyonis 3 Medway and others (1989).

Indiana B. procyonis 2 Fitzgerald and others (1991).

New York B. procyonis 2 W.B. Stone, written commun. (1998).

Massachusetts B. procyonis 3

2 F.S. Tseng, unpub. data (2007).

Washington B. procyonis 1 J.R. Huckabee, unpub. data (2001).

Western Canadian 

porcupine

Manitoba B. procyonis 1 Thompson and others (2008).

Brazilian porcupine Florida B. procyonis 1 R.L. Ball, written commun. (2000).

Nutria Michigan B. procyonis or 

B. columnaris4

20 of 35 Dade and others (1977).

Germany B. procyonis 65 Koch and Rapp (1981).

Mountain beaver Washington B. procyonis 3 J.R. Huckabee, unpub. data (2010).

North American beaver Ireland B. procyonis >3 Kelly and Innes (1966).

New York B. procyonis 2 W.B. Stone, written commun. (1998).

Ontario B. procyonis 3

1 B.J. McEwen, unpub. data (1990).

Ontario B. procyonis 3

1 D.J. Russell, unpub. data (2004).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Positive for Baylisascaris neural larva migrans, but without clinical central nervous system disease.

3

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae not found or animals survived.

4

 In some cases, involvement of B. columnaris could not be ruled out. However, most of these cases are most likely B. procyonis.
Species Susceptibility 47

Table 11. Rabbits with naturally occurring Baylisascaris neural larva migrans.1

[NR, not reported]

Host Location Probable parasite 

species

Number of 

animals 

affected

Reference

Eastern cottontail Illinois B. procyonis or B. columnaris3 1 Ferris and others (1960).

Virginia B. procyonis 16 of 60 Nettles and others (1975).

Connecticut B. procyonis or B. columnaris3 1 Church and others (1975).

Virginia B. procyonis 18 of 72 Jacobson and others (1976).

Iowa B. procyonis or B. columnaris3 NR Greve (1985).

Indiana B. procyonis 2

1 K.R. Kazacos, unpub. data (1978).

Indiana/Illinois B. procyonis 3 K.R. Kazacos, unpub. data (1984).

Illinois B. procyonis or B. columnaris3 1 R.H. Evans, unpub. data (1983).

Illinois B. procyonis 1 A.L. Shima and K.R. Kazacos, 

unpub. data (1985).

Ontario B. procyonis 1 I.K. Barker, unpub. data (1990).

Desert cottontail California B. procyonis 6 of 39 Evans (2002).

California B. procyonis 2

2 C.E. Stringfield and others, unpub. data (1995).

European rabbit 

(domesticated)

Connecticut B. procyonis Several Church and others (1975).

Michigan B. procyonis or

B. columnaris3

80 Dade and others (1975).

Indiana B. procyonis 20–25 Kazacos and others (1983).

Iowa B. procyonis NR Greve (1985).

Indiana B. procyonis 3 Boyce and others (1988a).

Indiana B. procyonis 15 Kazacos and Kazacos (1988).

Ontario B. procyonis NR Sanford (1989).

Washington B. procyonis 4 Deeb and DiGiacomo (1994).

Japan B. procyonis 36 Furuoka and others (2003).

Indiana B. procyonis Several H.L. Shivaprasad, unpub. data (1979).

Indiana B. procyonis 2 D.W. Knapp and K.R. Kazacos, 

unpub. data (1985).

Illinois B. procyonis 2 K.R. Kazacos, unpub. data (1996).

Illinois B. procyonis 6 L.J. Hardy and K.R. Kazacos, unpub. data (1998).

Illinois B. procyonis 4 P.J. Didier, unpub. data (1982).

Illinois B. procyonis 2 R.H. Evans, unpub. data (1983).

Indiana B. procyonis 6 K.R. Kazacos, unpub. data (1984).

Indiana B. procyonis 2

2 K.R. Kazacos, unpub. data (1986).

Indiana B. procyonis 3 N.A.Q. Mehdi, unpub. data (1982).

Indiana B. procyonis 20 D.D. Harrington, unpub. data (1982).

Indiana B. procyonis 1 H.L. Thacker, unpub. data (1984).

Indiana B. procyonis 1 J.A. Engelhardt, unpub. data (1987).

New York B. procyonis 3 L. Roth, written commun. (1983).

New York B. procyonis 2 W.B. Stone and K.R. Kazacos, 

unpub. data (1987).

Washington B. procyonis 2 M.M. Garner, written commun. (2010).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae not found or animals survived.

3

 In some cases, involvement of B. columnaris could not be ruled out; however, most of these cases are most likely B. procyonis.
48 Baylisascaris Larva Migrans

Table 12. Carnivores with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis]

Host Location Number of 

animals affected Reference

Red (silver) fox Iowa 4 of 4 Larson and Greve (1983); Greve (1985).

Domestic dog Michigan 1 Thomas (1988).

Indiana 1 Rudmann and others (1996).

California 1 Windsor and others (2009).

California 1 H.R. Galano and others, unpub. data (2005).

American badger California 1 Evans (2002).

Southern sea otter California 1 N.J. Thomas and others, unpub. data (1998).

Long-tailed weasel California 1 of 7 Evans (2002).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.
Species Susceptibility 49

Table 13. Primates with naturally occurring Baylisascaris neural larva migrans.1

Host Location

Probable 

parasite 

species

Number of 

animals 

affected

Reference

Black-and-white ruffed lemur Oklahoma B. procyonis 2 Campbell and others (1997).

Rhode Island B. procyonis 6 J.C. Martin,unpub. data (1998–99).

Tennessee B. procyonis 3 S.J. Barrett, unpub. data (1996).

California B. procyonis 1 M.M. Garner, written commun. (2010).

Missouri B. procyonis 2

7 of 11 A. Alexander and others, unpub. data (2013).

Red ruffed lemur Rhode Island B. procyonis 3 J.C. Martin, unpub. data (1998–99).

Ring-tailed lemur Ontario B. procyonis 3

1 T. Van Dreumel, unpub. data (2014).

White-headed lemur Spain B. procyonis 2 Jimenez Martinez and others (2015).

Coquerel’s giant 

mouse lemur 

California B. procyonis 1 K.R. Kazacos and F.H. Dunker, unpub. data (1995).

Mohol bushbaby Wisconsin B. procyonis 2

3 V.L. Clyde and R.S. Wallace, written 

commun. (2011).

White-headed marmoset Texas B. columnaris 3 Huntress and Spraker (1985); T.R. Spraker, 

written commun. (1986).

Illinois and Texas B. columnaris 1 K.R. Kazacos and P.L. Wolff, unpub. data (1986).

Black-mantled tamarin Texas B. columnaris 1 Huntress and Spraker (1985); T.R. Spraker, 

written commun. (1986).

Red-handed (Midas) tamarin Texas B. columnaris 1 Huntress and Spraker (1985); T.R. Spraker, 

written commun. (1986).

Indiana B. procyonis 1 M.C. Spriggs, unpub. data (2009).

Cottontop tamarin Louisiana B. procyonis 2

1 M.M. Garner, written commun. (2010).

Emperor tamarin California B. procyonis 1 M.M. Garner, written commun. (2010).

Golden-headed lion tamarin Maryland B. procyonis 2 Pessier and others (1997).

California B. procyonis 3 Stringfield and Sedgwick (1997); Pessier and 

others (1997).

Mantled guereza 

(Abyssinian black-andwhite Colobus monkey)

Indiana B. procyonis 1 J.A. Beck, written commun. (2014).

White-eared titi monkey Maryland B. procyonis 1 and 2

1 Beck and others (2010).

White-handed gibbon Kansas B. procyonis 1 Ball and others (1998).

Spider monkey Maryland B. procyonis 1 Garlick and others (1996).

De Brazza’s monkey Indiana B. procyonis 2 C.L. Eng and K.R. Kazacos, unpub. data (1997).

Japanese macaque Japan B. procyonis 9 Sato and others (2005).

Rhesus macaque Maryland B. procyonis 7 of 13 Gozalo and others (2008).

Crab-eating macaque Connecticut B. procyonis 1 Shoieb and Radi (2014).

Bornean orangutan Tennessee B. procyonis 1 D. Zimmerman and others, unpub. data (2005).

Bornean orangutan hybrid Wisconsin B. procyonis 1 Hanley and others (2006).

Human3 Pennsylvania B. procyonis 1 Huff and others (1984).

Illinois B. procyonis 1 Fox and others (1985).

California B. procyonis 1 Rowley and others (2000).

B. procyonis 1 Kazacos and others (2002).

Missouri B. procyonis 1 Moore (2009); Ciarlini and others (2011).

British Columbia B. procyonis 1 Hung and others (2012).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, with information and data on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae not found or animals survived.

3

 Does not include serologically diagnosed cases.
50 Baylisascaris Larva Migrans

Table 14. Miscellaneous mammals with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis]

Host Location Number of 

animals affected Reference

Marsupials

Red kangaroo Michigan 11 of 20 Agnew and others (1994).

Yellow-footed rock wallaby California 2

2 Stringfield and Sedgwick (1997).

Woylie California 2

1 J.E. Wynne, written commun. (1998).

Long-nosed potoroo New York 2

1 K.A. Volle, unpub. data (2010).

Bennett’s wallaby Indiana 1 J.A. Beck and others, unpub. data (2014).

Bats

Indian flying fox California 2

1 Stringfield and Sedgwick (1997).

Ontario 2

1 S.J. Best, unpub. data (1990).

Rodrigues flying fox Washington 2 K.E. Helmick, unpub. data (2007).

Ungulates

Domestic sheep Idaho 1 of 3 Anderson (1999).

Domestic cow Alberta 1 E. Janzen and others, unpub. data (2010).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.

Table 15. Gamebirds with naturally occurring Baylisascaris neural larva migrans.1

[NR, not reported]

Host Location Probable parasite 

species

Number of

 animals affected Reference

Domestic chicken Indiana B. procyonis 622 Richardson and others (1980).

Ontario B. procyonis 2

2 D.A. Smith, written commun. (2015).

Chukar Maryland B. procyonis 1 of 30 Sass and Gorgacz (1978).

Illinois B. procyonis 2

1 P.J. Didier, unpub. data (1984).

Ontario B. procyonis 8 D.A. Smith, written commun. (2015).

Rock partridge Ontario B. procyonis 2

1 I.K. Barker, unpub. data (2007).

Ruffed grouse New York B. procyonis 3 W.B. Stone, written commun. (1998).

Common pheasant Wisconsin B. procyonis 200–400 Kazacos and others (1986).

Wild turkey New York B. procyonis 2 W.B. Stone, written commun. (1998). 

Indian peafowl Nebraska B. procyonis 2

1 Armstrong and others (1989).

Ontario B. procyonis 1 and 21 D.A. Smith, unpub. data (1998; 2008).

Helmeted guineafowl Ontario B. procyonis 2

1 D.A. Smith, written commun. (2015).

Northern bobwhite Indiana B. procyonis 85 of 85 Reed and others (1981).

Iowa B. procyonis NR Greve (1985).

Kansas B. procyonis or

B. columnaris3

1 Williams and others (1997).

Illinois B. procyonis 640 of 712 K.R. Kazacos and others, unpub. data (2002). 

California quail California B. procyonis 7 of 29 Evans (2002).

Australian brushturkey Indiana and Missouri B. procyonis 1 Kazacos and others (1982).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.

3

 In some cases, involvement of B. columnaris could not be ruled out; however, most of these cases are most likely B. procyonis.
Species Susceptibility 51

Table 16. Perching birds with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis]

Host Location Number of 

animals affected Reference

House sparrow California 3 of 111 Evans (2002).

Bushtit California 3 of 3 Evans (2002).

Canary California 3 B.C. Barr, written commun. (1998).

California 12 of 22 Loretti and others (2008).

House finch California 8 of 56 Evans (2002).

Northern cardinal Indiana 1 K.R. Kazacos, unpub. data (2001).

Spotted towhee California 1 Evans (2002).

Loggerhead shrike California 1 of 13 Evans (2002).

California thrasher California 3 of 8 Evans (2002).

Northern mockingbird California 8 of 74 Evans (2002).

American robin Illinois 1 Evans and Tangredi (1985).

European starling California 3 of 68 Evans (2002).

Blue jay Illinois 2 Evans and Tangredi (1985).

Western scrub jay California 7 of 8 Evans (2002).

 American crow New York 1 B.P. Tangredi and K.R. Kazacos, unpub. data (1998).

California 1 of 122 Evans (2002).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

Table 17. Wading birds, shorebirds, diving birds, and waterfowl with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis]

Host Location Number of 

animals affected Reference

Black-crowned night heron California 2 of 38 Evans (2002).

Sanderling California 1 of 4 Evans (2002).

Inca tern California 2

1 M.M. Garner, written commun. (2010).

Mallard X California 2 of 3 Evans (2002).

Crested screamer Manitoba 2 Thompson and others, (2008).

Tennessee 2 D. Zimmerman and others, unpub. data (2003).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.
52 Baylisascaris Larva Migrans

Table 18. Parrots with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis]

Host Location Number of 

animals affected Reference

Budgerigar California 11 B.C. Barr, written commun. (1998).

Ontario 2

3 Russell (2006).

Eastern rosella Ontario 2

1 D.A. Smith, written commun. (2015).

Rosella, unknown species Ontario 2

4 D.A. Smith, written commun. (2015).

Blue-and-yellow macaw Nebraska 3 of 4 Armstrong and others (1987; 1989).

Scarlet macaw Nebraska 3 of 4 Armstrong and others (1987; 1989).

Iowa 2 M.A. Nieves and others, unpub. data (1989).

Red-and-green macaw California 1 M.M. Garner, written commun. (2010).

Hybrid blue-and-yellow 

macaw X scarlet macaw

Nebraska 2 of 2 Armstrong and others (1987; 1989).

Military macaw California 1 M.M. Garner, written commun. (2010).

African grey parrot Ontario 1 D.A. Smith, written commun. (2015).

Burrowing parrot Manitoba 4 of 5 Thompson and others (2008).

Blue-fronted Amazon California 1 B.C. Barr, written commun. (1998).

Yellow-headed Amazon Indiana 2 A.M. Lennox and others, unpub. data (1996); Lennox and others (2015).

Ontario 3 and 2

1 Russell (2006); D.A. Smith, written commun. (2015).

Yellow-naped Amazon Ontario 1 Russell (2006); D.A. Smith, written commun. (2015).

Cuban Amazon Ontario 1 Russell (2006); D.A. Smith, written commun. (2015).

Red-crowned Amazon California 1 Done and Tamura (2014).

Blue-crowned parakeet Indiana 5 A.M. Lennox and others, unpub. data (1996); Lennox and others (2015).

Orange-fronted parakeet Indiana 1 A.M. Lennox and others, unpub. data (1996); Lennox and others (2015).

Sun parakeet Indiana 2 A.M. Lennox and others, unpub. data (1996); Lennox and others (2015).

Rosy-faced lovebird Ontario 2

12 of 12 Russell and others (2005); Russell (2006).

Ontario 5 of 10 D.A. Smith, written commun. (2015).

Lovebird California 2

1 B.C. Barr, written commun. (1998).

Thick-billed parrot California 2

1 Stringfield and Sedgwick (1997).

California 1 C.L. Eng, unpub. data (2012).

Rainbow lorikeet Ontario 15 Russell and others (2005); Russell (2006); 

D.A Smith, written commun. (2015).

Swainson’s lorikeet Iowa 2

2 M.M. Garner, written commun. (2010).

Ontario 4 Russell and others (2005); Russell (2006).

Marigold (Edward’s) lorikeet Ontario 2

1 Russell (2006).

Yellow-backed lorikeet Ontario 2

1 I.K. Barker, unpub. data (2005).

Ornate lorikeet Ontario 2

1 D.A. Smith, written commun. (2015).

Lorikeet, unidentified species Missouri 2

36 A. Alexander and others, unpub. data (2013).

Cockatiel Iowa 3 of 3 Myers and others (1983); Greve (1985).

California 2

2 B.C. Barr, written commun. (1998).

California 34 of 35 Diab and others (2012).

Ontario 2

1 D.A. Smith, written commun. (2015).

Galah California 3 Stringfield and Sedgwick (1997).

Red-tailed black cockatoo California 2

1 Stringfield and Sedgwick (1997)

Salmon-crested cockatoo Kansas 1 Wolf and others (2007).

White (umbrella) cockatoo Ontario 2

1 D.A. Smith, written commun. (2015).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.
Species Susceptibility 53

Table 19. Flightless birds with naturally occurring Baylisascaris neural larva migrans.1

[>, greater than]

Host Location Probable 

parasite species

Number of 

animals affected Reference

Emu Indiana B. columnaris 2 Winterfield and Thacker (1978); Kazacos, 

Winterfield and others (1982).

Indiana B. procyonis 2

2 Kazacos and others (1991).

Indiana B. procyonis 2

2 K.R. Kazacos, unpub. data (1996).

Ontario B. procyonis 9 chicks

2

2 adults

Smith and others (1993); Kwiecien and others 

(1993).

Ontario B. procyonis 2

1 I.K. Barker, unpub. data (1995).

Missouri B. procyonis 2 of 2 K.R. Kazacos, unpub. data (1994).

Kansas B. procyonis 2 of 4 Suedmeyer and others (1996).

Oklahoma B. procyonis 2

3 Campbell and others (1997).

California B. procyonis 5 of 9 Loretti and others (2008).

Michigan B. procyonis 1 D.W. Agnew and K.R. Kazacos, unpub. data (1996).

Kansas, Missouri, 

Nebraska, Indiana, 

New York, 2

California

B. procyonis >18 K.R. Kazacos, unpub. data (1985–96).

California B. procyonis 2

1 F.H. Dunker and K.R. Kazacos, unpub. data (1997).

California B. procyonis 1 B.C. Barr, written commun. (1998).

California B. procyonis 6 L.W. Woods, written commun. (2015).

Indiana B. procyonis 2

1 M.C. Spriggs, unpub. data (2009).

Ostrich Indiana B. procyonis 1 Kazacos and others (1991).

Ontario B. procyonis 2 and 2

2 D.A. Smith, written commun. (2015).

Greater rhea Indiana B. procyonis 2

2 K.R. Kazacos, unpub. data (1996).

Pennsylvania B. procyonis 1 M.M. Garner, written commun. (2010).

Ontario B. procyonis 2 D.A. Smith, written commun. (2015).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.
54 Baylisascaris Larva Migrans

Table 20. Miscellaneous birds with naturally occurring Baylisascaris neural larva migrans.1

[Probable parasite species B. procyonis; >, greater than]

Host Location Number of 

animals affected Reference

Rock pigeon Oregon 10 of 60 Helfer and Dickinson (1976).

Illinois 1 Evans and Tangredi (1985).

British Columbia 2 Coates and others (1995).

Nebraska >15 of 200 E.N. Pendleton and V.W. Rinne, unpub. data (1992).

Ontario 6 D.A. Smith, written commun. (2015).

Mourning dove New York 2 Evans and Tangredi (1985).

Illinois >25 C.U. Meteyer and others, unpub. data (1992).

Missouri 1 R.H. Evans, written commun. (1983).

California 30 of 81 Evans (2002).

Diamond dove California 2

1 B.C. Barr, written commun. (1998).

Northern flicker Washington 3 J.R. Huckabee, unpub. data (2010).

Greater roadrunner California 3 of 21 Evans (2002).

Northern red-billed hornbill Ontario 2

1 D.A. Smith, written commun. (2015).

Speckled mousebird Ontario 5 D.A. Smith, unpub. data (2006); I.K. Barker, 

unpub. data (2006).

Blue-naped mousebird Ontario 2

1 I.K. Barker, unpub. data (2006).

Barn owl California 1 Evans (2002).

1

 Cases have been confirmed by the author or cooperating board-certified pathologists, and information and data are on file with the author.

2

 Probable case based on characteristic lesions, clinical signs, and (or) history of exposure; larvae were not found or animals survived.
Species Susceptibility 55

Table 21. Animals experimentally susceptible to Baylisascaris neural larva migrans.1—Continued

[NR, not reported]

Animal Location Parasite

Number of animals 

affected per total 

number of animals

Reference

Rodents

House mouse Illinois B. procyonis 3 of 3 Tiner (1952a).

Laboratory mouse Illinois B. procyonis 12 of 12; 15 of 21; 

8 of 9 

Tiner (1949; 1952a; 1953a).

Illinois B. columnaris 3 of 9; 7 of 11 Tiner (1952a; 1953a).

Australia B. columnaris 4 of 9; 12 of 32; 

2

2 of 9; 21 of 5

Sprent (1953a; 1955).

Bulgaria B. transfuga 2

16 of 20 Matoff and Komandarev (1965).

Minnesota B. columnaris 54 of 70 Clark and others (1969).

Indiana B. columnaris 9 of 31 K.R. Kazacos, unpub. data (1984).

Indiana B. columnaris 7 of 22 K.R. Kazacos, unpub. data (1985).

Indiana B. procyonis 12 of 12 K.R. Kazacos, unpub. data (1980).

Indiana B. procyonis 16 of 16 K.R. Kazacos, unpub. data (1985).

Indiana B. columnaris 0 of 12 Boyce and others (1988b).

Kansas B. procyonis 6 of 6 Lindquist (1978).

Kansas B. procyonis NR Al-Lebban and Lindquist (1979).

Indiana B. procyonis 25 of 25 Kazacos (1981).

Ohio B. procyonis 66 of 66 Dubey (1982).

Indiana B. procyonis 10 of 10 Wirtz (1982).

Indiana B. procyonis 215 of 215 K.R. Kazacos, unpub. data (1984).

Indiana B. melis 12 of 12 K.R. Kazacos, unpub. data (1986).

Indiana B. procyonis 12 of 12 Boyce and others (1988b).

Indiana B. procyonis 5 of 5 K.R. Kazacos, unpub. data (1989).

Japan B. procyonis 70 of 75 Miyashita (1993).

Italy B. transfuga 5 of 80 Papini and Casarosa (1994).

Indiana B. procyonis 24 of 25 Garrison (1996).

Indiana B. procyonis 28 of 30 Sheppard and Kazacos (1997).

Japan B. procyonis 36 of 43 Sato and others (2004).

Japan B. transfuga 4 of 15 Sato and others (2004).

Laboratory rat New Jersey B. procyonis 2

2 of 3 Tiner (1954).

Indiana B. procyonis 4 of 36 K.R. Kazacos, unpub. data (1982).

Indiana B. procyonis 12 of 19 Wirtz (1982).

Indiana B. columnaris 0 of 48 K.R. Kazacos, unpub. data (1984).

Mongolian gerbil Japan B. procyonis 13 of 15 Akao and others (2003).

Japan B. procyonis 39 of 41 Sato and others (2004).

Japan B. transfuga 13 of 13 Sato and others (2004).

White-footed mouse Illinois B. procyonis NR; 7 of 10 Tiner (1949; 1953a).

Illinois B. columnaris 0 of 6; 1 of 4 Tiner (1953a).

Indiana B. procyonis 17 of 30 Sheppard and Kazacos (1997).

Western harvest 

mouse

Indiana B. procyonis 3 of 3 Sheppard (1996).
56 Baylisascaris Larva Migrans

Table 21. Animals experimentally susceptible to Baylisascaris neural larva migrans.1—Continued

[NR, not reported]

Animal Location Parasite

Number of animals 

affected per total 

number of animals

Reference

Rodents—Continued

Meadow vole Indiana B. procyonis 29 of 30 Sheppard (1996).

Ontario B. columnaris 2

8 Berry (1985).

Prairie vole Indiana B. procyonis 3 of 4 Sheppard (1996).

Golden hamster Illinois B. procyonis NR Tiner (1949).

Indiana B. columnaris 5 of 8 K.R. Kazacos, unpub. data (1984).

Indiana B. columnaris 0 of 14 K.R. Kazacos, unpub. data (1985).

Indiana B. procyonis 32 of 32 Kazacos (1981).

Indiana B. procyonis 23 of 23 Wirtz (1982).

Indiana B. procyonis 54 of 54 K.R. Kazacos, unpub. data (1984).

Allegheny woodrat Indiana B. procyonis 26 of 26 K.R. Kazacos, unpub. data (1997).

Hispid cotton rat Illinois B. procyonis 6 of 6; 7 of 8; 3 of 3 Tiner (1949, 1952a; 1953a)

Illinois B. columnaris 0 of 4 Tiner (1952a).

Meadow jumping 

mouse

Indiana B. procyonis 1 of 1 Sheppard (1996).

Eastern chipmunk Indiana B. procyonis 1 K.R. Kazacos, unpub. data (1981).

Eastern gray squirrel Illinois B. procyonis 4 of 6 Tiner (1949; 1952a; 1953a).

Indiana B. procyonis 10 of 10 Wirtz (1982).

Indiana B. procyonis 20 of 20 K.R. Kazacos, E.A. Kazacos, and 

Vestre (1984)

Indiana B. columnaris 0 of 1 K.R. Kazacos, unpub. data (1984).

Woodchuck Connecticut B. columnaris 1 of 5; 2

4 of 5 Swerczek and Helmboldt (1970).

Indiana B. procyonis 5 of 5 K.R. Kazacos, unpub. data (1981–83).

Domestic guinea pig Illinois B. procyonis NR Tiner (1949; 1953b).

Pennsylvania B. procyonis NR Donnelly and others (1989).

Rabbits and hares

Eastern cottontail Illinois B. procyonis NR Tiner (1954).

Virginia B. procyonis 1 of 1 Jacobson and others (1976).

Domestic rabbit Illinois B. procyonis NR Tiner (1954).

Connecticut B. columnaris 3 of 4 Church and others (1975).

Indiana B. procyonis 18 of 18 K.R. Kazacos, unpub. data (1980).

Indiana B. procyonis 1 of 1 Boyce and others (1989).

Shrews

Northern short-tailed 

shrew

Indiana B. procyonis 0 of 9 Sheppard (1996).

Carnivores

Domestic dog Illinois B. procyonis 3 of 5 Snyder (1983).

Indiana B. procyonis 0 of 8 K.R. Kazacos, unpub. data (1981).

Japan B. procyonis 0 of 3 Miyashita (1993).

Domestic cat Indiana B. procyonis 0 of 13 K.R. Kazacos, unpub. data (1981).

Japan B. procyonis 0 of 1 Miyashita (1993).
Species Susceptibility 57

Infecting Dose and Onset of Disease

B. procyonis is not a parasite that must migrate within 

or through the nervous system of host animals (that is, it is 

not neurotropic), and it is estimated that only 5–7 percent of 

ingested larvae enter the brains of paratenic hosts. Similar to 

humans, animals with low-level infection will remain asymptomatic if no larvae enter the brain (Kazacos, 2001); however, 

a single B. procyonis larva in the brain of a small mammal or 

bird is usually fatal (Tiner, 1953a,b; Sheppard and Kazacos, 

1997; K.R. Kazacos, unpub. data, 1980–85). In natural cases 

of clinical NLM, one to five or more larvae are often recovered from the brain during postmortem examination of an 

animal, and in some cases hundreds to several thousand larvae 

were present in the brain, due to heavy infection with eggs 

(Tiner, 1953a; Fox and others, 1985; Armstrong and others, 

1989; Van Andel and others, 1995; Lennox and others, 2015). 

Those larvae migrating in the brain produce traumatic damage 

and inflammation, resulting in progressive neurologic disease, 

the onset and severity of which are related to the number of 

larvae. Animals that are infected with high numbers of larvae 

may become dull and lethargic and stop eating. Labored 

breathing and increased respiratory rates develop 2–5 days 

following infection (postinfection), due to hemorrhage and 

inflammation in the lungs caused by larval migration through 

the lungs (Church and others, 1975; Kazacos, Wirtz, and 

Table 21. Animals experimentally susceptible to Baylisascaris neural larva migrans.1—Continued

[NR, not reported]

Animal Location Parasite

Number of animals 

affected per total 

number of animals

Reference

Carnivores—Continued

Domestic ferret Indiana B. procyonis 4 of 4 Kazacos (1981); E.A. Kazacos and 

K.R. Kazacos (1988).

Least weasel Indiana B. procyonis 1 K.R. Kazacos, unpub. data (1982).

Primates

Squirrel monkey Indiana B. procyonis 4 of 4 Kazacos, Wirtz and others (1981)

Japan B. potosis 0 of 2 Tokiwa and others (2015).

Crab-eating macaque Indiana B. procyonis 4 of 4 Kazacos, Vestre and Kazacos (1984).

Olive baboon Georgia B. procyonis 1 M.L. Eberhard and others, unpub. data (2002).

Ungulates

Domestic pig Indiana B. procyonis 0 of 6 K.R. Kazacos and E.A. Kazacos (1984).

Birds

Domestic chicken Indiana B. procyonis 17 of 50 Kazacos and Wirtz (1983).

Italy B. transfuga 2

32 of 35 Papini and others (1993).

Cockatiel Ontario B. procyonis 24 of 27 Russell (2006).

Domestic duck Indiana B. procyonis 8 of 21 Wirtz (1982).

1

 Information and data are on file with the author.

2

 Positive for Baylisascaris neural larva migrans, but without clinical central nervous system disease.

others, 1981; Wyand-Ouellette and others, 1983; Donnelly 

and others, 1989; Kazacos, 1997). Although larvae enter the 

somatic tissues, eyes, and brain of laboratory rodents as early 

as 3 days postinfection (Tiner, 1953b; Kazacos and others, 

1985; Kazacos, 1986), clinical CNS disease is not usually 

apparent before 9–10 days postinfection, due to the lag time 

in development of CNS damage and inflammation (Kazacos, 

1997; Sheppard and Kazacos, 1997). Infection among animal 

species also varies, based on apparent differences in larval 

migration patterns and host responses; for example, the onset 

of clinical signs was 20–21 days postinfection, on average, 

in white-footed mice compared with 9–14 days postinfection 

in white laboratory mice, and fewer larvae were recovered 

from the anterior carcass, head, and brain and more from the 

abdominal and thoracic viscera of white-footed mice (Sheppard, 1995; Sheppard and Kazacos, 1997). Nonetheless, single 

larvae entering the brains of white-footed mice produced fatal 

NLM. In larger animals, signs may not become apparent until 

at least 2–4 weeks postinfection, which is related to the animals’ larger brain sizes and the greater infective dose needed 

to cause clinical disease. If the larvae leave the brain proper 

and (or) become encapsulated, clinical signs may abate or 

stabilize and the animal can survive and function, albeit with 

variable CNS deficits depending on the location and severity 

of migration damage.
58 Baylisascaris Larva Migrans

Clinical Signs

Clinical signs in rodents and other small mammals are 

directly related to systemic migration and CNS damage (table 

22; fig. 9; fig. 10; fig. 11). A 12-week-old beagle puppy was 

lethargic, had progressive rear-limb weakness, and was unable 

to swallow, circled, fell onto its side, and was apparently blind. 

Its condition worsened rapidly; it developed a severe head tilt, 

lay on its side, became stuporous, and paddled its legs aimlessly (Rudmann and others, 1996). Similar signs were seen 

in experimentally infected squirrel monkeys (Kazacos, 2001; 

Kazacos, Wirtz, and others, 1981), and matched those seen 

in naturally infected nonhuman primates and children with 

Baylisascaris (Huff and others, 1984; Fox and others, 1985; 

Huntress and Spraker, 1985; Kazacos, 1996b, 2000, 2001; 

Campbell and others, 1997; Ball and others, 1998). Mildly 

affected nonhuman primates showed only subtle forelimb 

tremors or stumbling (C.L. Eng and K.R. Kazacos, unpub. 

data, 1997) or had slight head tilts and incoordination (ataxia)

(Pessier and others, 1997), indicating low-level and (or) more 

covert infection, a possibility that should be considered more 

often in the diagnosis of humans with this infection.

Table 22. Clinical signs of Baylisascaris neural larva migrans 

in small mammals.

[Compiled from Church and others (1975); Kazacos (2001); Kazacos and 

Boyce (1989); Sheppard and Kazacos (1997); Wirtz (1982); and various 

case reports]

Initial clinical signs

Decreased activity.

Depression or nervousness.

Rough hair coat.

Tremors in the front paws.

Slight head and (or) body tilts.

Circling or jumping when disturbed.

Worsening clinical signs; various combinations of:

Severe head and (or) body tilts.

Slow arching of the head and neck with “stargazing.”

Extension and rigidity of the forelimbs.

Motor weakness.

Incoordination (ataxia).

Continuous circling.

Leaning and (or) falling over.

Arching of the body.

Lying on the side (lateral recumbency).

Paddling movements while lying down.

Rolling.

Blindness.

Rapid eye movements.

Coma.

Death.

Figure 9. Mammals affected with Baylisascaris procyonis neural 

larva migrans. 

Ferret with arching of 

the body and head and 

extension of its forelimbs. 

(Photo by Kevin Kazacos 

and Sam Royer)

Squirrel monkey lying prostrate 

and semicomatose. (Photo by 

Kevin Kazacos and Sam Royer) 

Woodchuck lying on its 

side and paddling its 

forelimbs. (Photo by Kevin 

Kazacos and Sam Royer)

Gray squirrel with severe 

head tilt and extension of its 

forelimbs. (Photo by Kevin 

Kazacos and Sam Royer)

Domestic rabbit with arching 

of the head and neck. (Photo 

by Kevin Kazacos and Sam 

Royer)

Domestic rabbit with arching 

of the head and neck and 

extension of its forelimb. 

(Photo by Kevin Kazacos 

and Sam Royer)

White-footed mouse recumbent 

on its side with head tilt and 

rigidity of its forelimbs. (Photo 

by Claudia Sheppard)

Chipmunk with head tilt. (Photo 

by Kevin Kazacos and Sam 

Royer)
Figure 10. The sequence of clinical neural larva migrans in a wild gray squirrel. A, B, C, The slow arching of the head and neck with 

“stargazing”’ progressing to leaning of the body. D, The squirrel with a wide stance, trying to steady itself while eating a sunflower seed. 

E, F, The squirrel has fallen over, lying on its side while continuing to eat the seed. (Photos by Kevin Kazacos)

D E F

Figure 11. Abnormal circling behavior of a mouse or vole (dead in photo) in the snow along the Bighorn River near Worland, 

Wyoming, likely caused by Baylisascaris neural larva migrans. (Photo by David G. Frahm)

A B C

Species Susceptibility 59
60 Baylisascaris Larva Migrans

Clinical signs in birds are similar to those seen in mammals (table 23; fig. 12). Clinically affected ostriches and emus 

show varying degrees of incoordination, with loss of equilibrium and balance, muscle weakness, wobbling, and progressive ataxia. An infected ostrich typically staggers, walks in 

circles, assumes a wide, splay-legged stance with its head 

extended downward for balance, and walks rapidly backward, 

stumbling and falling; eventually it is unable to stand or walk 

and becomes increasingly emaciated (Kazacos, Fitzgerald, 

and Reed, 1991; Kwiecien and others, 1993; Suedmeyer and 

others, 1996). It is quite impressive that this degree of clinical 

disease in a large bird such as an ostrich can be caused by one 

or two Baylisascaris larvae in the brain; however, despite its 

large body size, an ostrich’s brain is relatively small and suffers considerable damage.

Table 23. Clinical signs of Baylisascaris neural larva migrans 

in birds.

[Compiled from Coates and others (1995); Kazacos ( 2001); Kazacos and 

Wirtz (1983); Kazacos and others (1991); Kwiecien and others (1993); 

A. M. Lennox and others, unpub. data (1996); Richardson and others 

(1980); Reed and others (1981); Russell (2006); Russell and others (2005); 

Wirtz (1982)]

Various combinations of:

Ruffled feathers.

Disorientation.

Head tremors or bobbing.

Twisted neck.

Inability to stand.

Muscle weakness and (or) tremors.

Rigidity of the legs.

Blindness.

Vocalizations.

Poor grip reflexes.

Incoordination.

Loss of balance.

Walking in circles.

Falling.

Rolling.

Inability to fly or loss of flight control.

Paralysis of one or both wings or legs.

Lying on the side with eyes closed.

Death.

Figure 12. Birds affected with Baylisascaris procyonis neural 

larva migrans.

Emu (with B. columnaris) 

unable to stand or use its 

legs. (Photo by Sam Royer)

Cardinal with twisted neck. 

(Photo by Kevin Kazacos and 

Sam Royer)

Bobwhite with twisted neck, 

ruffled feathers, and rigidity 

of the feet. (Photo by Kevin 

Kazacos and Andy Dziubinskyj)

Domestic chicken with 

twisted neck and rigidity of 

the legs. (Photo by Kevin 

Kazacos and Sam Royer)

Domestic duck with twisted 

neck and inability to stand. 

(Photo by Kevin Kazacos and 

Sam Royer)

Blue and gold macaw unable 

to stand or hold its head up. 

(Photo by Kevin Kazacos and 

Sam Royer)
Species Susceptibility 61

Granuloma Formation

Following migration in mammals, larvae become encapsulated in various tissues in well-circumscribed granulomas

(fig. 13). Larval granulomas are white, easily visible (1–3 

millimeters in diameter) and are found in many organs and 

tissues, including the skeletal muscles, liver, lungs, heart, diaphragm, pancreas, spleen, kidneys, abdominal lymph nodes, 

intestinal wall and connective tissue, brain, and eyes (Sprent, 

1952a; Kazacos, Wirtz, and others, 1981; Kazacos, 1986, 

1997, 2001; Sato and others, 2004). The development and (or) 

distribution of B. procyonis larval granulomas varies somewhat with animal species, but they are generally found in these 

locations in all types of infected mammals during necropsy 

examinations for this infection (Sheppard and Kazacos, 1997; 

Kazacos, 2001; Sato and others, 2004) (table 24).

On the other hand, gross lesions typically are not found 

during necropsies of birds, and tissue alterations are usually 

confined to the brain, especially within the cerebellum (Kazacos and Boyce, 1989; Kazacos, 2001). No larval granulomas 

were found grossly in experimentally infected chickens and 

ducks, and only a single granuloma was seen in an outer 

muscle of the eye during microscopic examination of tissue 

(Wirtz, 1982). Solitary larval granulomas were also found in 

the lungs of a naturally infected brushturkey and bobwhite 

(Kazacos, Kazacos, and others, 1982; Williams and others, 

Figure 13. Larval granulomas on the intestinal tract of a hamster with B. columnaris neural 

larva migrans. (Photo by Kevin Kazacos and Sam Royer)

1997). The comparative lack of gross lesions and scarcity of 

larval granulomas in birds with Baylisascaris NLM has been 

reported in numerous natural cases and outbreaks (Richardson 

and others, 1980; Reed and others, 1981; Myers and others, 

1983; Evans and Tangredi, 1985; Armstrong and others, 1989; 

Kazacos and others, 1991; Kwiecien and others, 1993; Russell 

and others, 2005; Russell, 2006; Lennox and others, 2015). In 

most clinical cases of NLM in birds, including large flightless birds, few or no larvae are found in somatic or visceral 

tissues, even though one to three larvae are found in the brain. 

Thus, most cases in birds appear to involve low-level infections and a higher probability of larval migration to the brain. 

However, much higher infection levels have occasionally been 

documented in the somatic tissues and viscera of birds. In an 

outbreak involving a mixed collection of parrots and conures, 

17–150 (mean, 87) B. procyonis larvae were recovered from 

the brains and 47–285 (mean, 173) larvae from the viscera 

and carcasses of 6 birds dying from rapid onset, severe CNS 

disease (Lennox and others, 2015); no gross lesions or larval 

granulomas were seen during necropsy. This difference in the 

reaction of birds, compared to mammals, may be related to 

differences in immune system recognition of larvae or their 

products and (or) mobilization of different types of inflammatory cells, including those involved in granuloma formation; 

it is well known that inflammation involving multiple types of 

cells in the brains of birds can be quite intense. 
62 Baylisascaris Larva Migrans

Table 24. Examples of locations of Baylisascaris larval granulomas in mammals.

Animal Body sites References

Mice and other 

small rodents

Heart surfaces and muscle, lungs, diaphragm, and 

skeletal muscles especially front end of the carcass, along the intestinal tract, other locations

Tiner (1953a,b); Kazacos (2001); Sheppard and 

Kazacos (1997); Wirtz (1982).

Gray squirrel Wall of the veins that return blood to the heart, the 

heart, lungs, diaphragm, intercostal muscles and 

other skeletal muscles, along the intestinal tract, 

other locations

Tiner (1953a); Kazacos (2001) and unpub. 

data (1984); Wirtz (1982).

Rabbit Heart, lungs, diaphragm, liver, mesentery (the 

membrane that envelops the intestines), along the 

intestinal tract, skeletal muscles, other locations

Church and others (1975); Kazacos (2001); 

Kazacos and others (1983); Nettles and others 

(1975); K.R. Kazacos, unpub. data (1985).

Monkey, lemur, marmoset Heart and great vessels, lungs, diaphragm, liver, 

stomach wall, mesentery, along intestinal tract, 

intercostal muscles, other skeletal muscles, 

other locations

Kazacos (2001); Kazacos, Wirtz and others (1981); 

K.R. Kazacos, unpub. data (1998); K.R. Kazacos 

and P.L. Wolff, unpub. data (1986).

Woodchuck, kangaroo Heart, lungs, liver, skeletal muscles; along the 

intestinal tract, especially the pouch at the beginning of the large intestine (cecum) and the colon

Richter and Kradel (1964); Jacobson and others 

(1976); Fleming and Caslick (1978); Agnew and 

others (1994); Swerczek and Helmboldt (1970).

Obtaining a Diagnosis

Diagnosis of Baylisascaris larva migrans in animals or 

humans is based on a combination of clinical, laboratory, and 

pathologic findings. Of these, only identification of larvae in 

or from the tissues is a confirmed diagnosis, although positive 

serology would indicate infection (Kazacos, 2001).

Unlike the situation in animals, relatively few clinical 

cases of Baylisascaris NLM in humans have been documented 

in North America and elsewhere, making physicians less likely 

to consider this dangerous pathogen in the diagnoses of their 

patients. Most documented human cases involved heavy infections and severe clinical signs, and it is likely that cases of 

lower level infection with more subtle or nonspecific signs are 

routinely missed. The disease consequences of B. procyonis

infection are so important that it simply must be considered in 

cases of neurologic disease in children as well as cases of eye 

disease in all age groups, especially those involving eosinophilic meningoencephalitis and DUSN, respectively.

Diagnosis of adult Baylisascaris infection in raccoons, 

skunks, dogs, kinkajous, or other species is based on identification of their eggs in the feces by using fecal flotation methods, identification of worms passed by the animal, or necropsy 

and examination of the small intestinal contents for the worms 

(Kazacos, 2001, 2006; Page and others, 2005). Young animals 

and occasionally others may be infected but not pass eggs in 

their feces, based on the presence of developing worms that 

are still immature and not yet shedding eggs; this is a so-called 

prepatent infection (Kazacos and Boyce, 1989). These animals 

still pose a threat to others, because as the worms mature, they 

will proceed to egg shedding in a few days to weeks, leading 

to contamination of the environment. Raccoons and probably 

other definitive hosts may also have intermittent and uneven 

egg shedding, related to the intensity of infection and (or) the 

age of worms present (Reed and others, 2012).
Obtaining a Diagnosis 63

Diagnosis of Baylisascaris Neural Larva Migrans

Diagnosis of Baylisascaris neural larva migrans 

(NLM) in animals or humans is based on a combination 

of clinical signs, a history of exposure to the parasite, and 

results of serology, imaging, and other diagnostic tests, 

as well as postmortem gross and histopathologic lesions. 

However, only identification of larvae in or from the tissues 

provides a confirmed diagnosis (Huff and others, 1984; 

Kazacos, Vestre, Kazacos, and Raymond, 1984; Kazacos, 

Vestre, and Kazacos, 1984; Kazacos, Raymond, and others, 

1985; Fox and others, 1985; Kazacos, 1991, 1997, 2000, 

2001; Goldberg and others, 1993; Murray and Kazacos, 

2004; Gavin and others, 2005).

Cerebrospinal Fluid Analysis

Diagnosis of Baylisascaris NLM in ill humans or 

animals depends on an assessment of clinical signs, compatible exposure history, serology, and evidence of white 

blood cells called eosinophils in the cerebrospinal fluid

(CSF), so-called eosinophilic pleocytosis, which often will 

accompany increased numbers of eosinophils in the blood 

(Kazacos, 2000, 2001; Murray and Kazacos, 2004; Gavin 

and others, 2005; Windsor and others, 2009; Kazacos and 

others, 2013). Eosinophil numbers appear to be higher in the 

CSF during acute disease, and their levels are related to the 

amount of damage and eosinophilic inflammation caused 

by the migrating larvae. In humans with severe NLM, so 

many eosinophils are in the central nervous system (CNS) 

that toxic eosinophil-derived proteins are released into brain 

tissue (Hamann and others, 1989). These proteins have been 

isolated directly from the CSF of affected children (Moertel 

and others, 2001), and they appear to be important contributors to the neurologic pathology and clinical signs associated with the infection. Similar proteins are likely released 

into the CNS of animals with NLM (Kazacos, 2001) and 

into the eye in cases of ocular larva migrans (OLM–DUSN) 

(Kazacos, 1996a).

Imaging Methods

Medical imaging techniques, including computed 

tomography (CT) and magnetic resonance imaging

(MRI), are useful for assessing the amount of brain damage and postinfection atrophy associated with Baylisascaris

and for locating lesions in particular areas of the brain or 

spinal cord in humans (Huff and others, 1984; Fox and 

others, 1985; Cunningham and others, 1994; Rowley and 

others, 2000; Kelly and others, 2009, 2012; Peters and 

others, 2012) and animals (Ball and others, 1998; Hanley 

and others, 2006). Baylisascaris NLM in children is usually 

associated with localized or widespread brain lesions deep 

in the white matter that are shown by MRI (A). MRI showed 

a large lesion in the frontal cortex of a gibbon (Ball and others, 1998) and involvement of deep central white matter in 

an orangutan with NLM (Hanley and others, 2006).

Serology

Serologic methods including indirect immunofluorescence, Western blotting, and enzyme-linked immunosorbent assay (ELISA), using culture-derived or recombinant DNA-generated larval excretory-secretory (ES)

antigens, are very useful for diagnosing Baylisascaris NLM 

in humans (Boyce and others, 1988a,b; 1989; Conraths and 

others, 1996; Moertel and others, 2001; Kazacos, 2001; 

Dangoudoubiyam and Kazacos, 2009; Dangoudoubiyam 

and others, 2010, 2011; Rascoe and others, 2013). The ES 

antigen ELISA has been used as a screening test, and even 

though it is known to cross-react with Toxocara spp. and 

some other parasites, the geographical distribution and epidemiology of most other causes of eosinophilic meningitis

(except for Toxocara spp.) do not overlap with Baylisascaris

NLM (Rowley and others, 2000). Further identification and 

(or) confirmation is done using the recombinant ES antigen 

ELISA and (or) Western blotting, which can separate Baylisascaris from Toxocara infection based on greater specificity and the recognition of specific ES proteins, respectively 

(Dangoudoubiyam and Kazacos, 2009; Dangoudoubiyam 

and others, 2010, 2011). The screening ELISA showed that 

8 percent of 389 nonclinical 1-to-4-year-old Chicago children were seropositive for Baylisascaris infection (Brinkman and others, 2003), as were 29 percent of 259 nonhuman 

primates at 6 major zoos (Zimmerman and others, 2010). 

The current “gold standard” for serologic diagnosis in 

humans involves Western blotting using the recombinant 

ES antigen (Dangoudoubiyam and others, 2010; Rascoe 

and others, 2013), which is highly sensitive and specific. 

Testing is available at the Centers for Disease Control and 

A. Magnetic resonance images 

of the brain of a child affected 

by Baylisascaris NLM, showing 

widespread deep white matter 

abnormalities. (Image by 

Howard A. Rowley) 
Diagnosis of Baylisascaris Neural Larva Migrans

64 Baylisascaris Larva Migrans

Prevention in Atlanta, Georgia, and the National Reference 

Centre for Parasitology in Montreal, Canada. By use of this 

assay, 7 of 43 nonclinical raccoon trappers from Georgia 

were seropositive for B. procyonis infection (C.A. Hall and 

others, unpub. data, oral commun., 2014). Better data on 

human seroprevalence will be obtained when the recombinant ES antigen Western blot is used to test blood samples 

from a wide segment of the population. Such methods could 

also be developed and applied to diagnosing NLM in various animal species.

Necropsy and Tissue Collection

Noncarnivorous animals suffering from neurologic 

disease should be captured if possible and submitted alive to 

a veterinary diagnostic laboratory with a request to examine 

them for Baylisascaris in addition to other causes. Dead animals should be submitted promptly, unless a field necropsy 

is done. 

For Baylisascaris NLM, because most larvae are 

encapsulated in granulomas in various parts of the body 

other than the brain, tissues should be searched for these and 

representative samples collected and preserved. Baylisascaris larvae can be dissected from these granulomas and 

identified microscopically, thereby aiding in the diagnosis of 

NLM (B, C).

Histopathology

Many diagnostic laboratories rely heavily on histopathology for disease diagnosis, but this has important 

limitations in the diagnosis of Baylisascaris larva migrans. 

Even in clinically affected animals, only a few (one to five) 

larvae may be present in the brain (Armstrong and others, 1989). In such cases, finding a larva in histopathologic 

B. Baylisascaris larval granulomas (arrows) on the heart 

of a woodchuck with NLM. (Photo by Kevin Kazacos 

and Sam Royer)

C. Larval granulomas on the intestinal tract of a marmoset 

with B. columnaris NLM. (Photo by Kevin Kazacos)

sections is a matter of chance, which will increase when 

either more larvae are present or more slides are examined 

(Kazacos, 1997, 2001). Often, however, histopathologic 

lesions compatible with Baylisascaris migration are seen, 

and a tentative or probable diagnosis is made in the absence 

of identified larvae.

Larval Isolation

Brain tissue can be processed in various ways to find 

larvae, including by squash techniques, Baermann funnel preparations, or digestion (Kazacos, 2001). Other 

tissues (heart, lung, etc.) can also be processed for larvae 

by digestion. These methods are particularly useful if 

multiple affected animals are available for examination. 

In the brain squash method, small pieces (approximately 

1 gram) of tissue are placed between two glass plates and 

pressed out until they are very thin (Kazacos, 2001) (D, 

E). The preparation is then examined under a dissecting 

microscope and searched for the presence of whole living 

larvae in the tissue (F, G), which are then recovered (H). 

Although it is time consuming for a large brain, this method 

works extremely well for small brains and will find single 

larvae if present. Brain tissue can also be finely minced and 

placed on cheesecloth in a Baermann funnel in warm saline 

or water, and larvae can be collected after a few hours or 

overnight (Richardson and others, 1980; Reed and others, 

1981; Fox and others, 1984; Kazacos, 2001). A drawback of 

this technique is that not all larvae may make it out of the 

tissue and down through the funnel, so in light infections 

they may be missed, but the method is particularly useful 

if multiple affected animals are examined. Finally, artificial 

digestion of blended or finely minced tissues by using a 

pepsin:hydrochloric acid mixture, followed by filtering and 

settling, is a very efficient method of larval recovery, especially for muscle tissue or viscera (Kazacos, 2001).
Obtaining a Diagnosis 65

D. Brain squash of a white-footed mouse with two Baylisascaris

larvae in the brain (circles). (Photo by Kevin Kazacos and Sam Royer)

H. B. procyonis larva recovered from brain squash of conure with 

NLM. (Photo by Kevin Kazacos and Sam Royer)

E. Brain squash of a conure with severe NLM, showing 49 larvae (circles) 

in about one-third of its brain. (Photo by Kevin Kazacos and Sam Royer)

F. Positive brain squash from a conure showing multiple larvae 

(arrows). (Photo by Kevin Kazacos and Sam Royer)

G. Higher magnification of conure brain squash showing B. procyonis

larva (arrow). (Photo by Kevin Kazacos and Sam Royer)

Diagnosis of Baylisascaris Neural Larva Migrans

Molecular Analysis

Recently, the complete sequence of DNA contained 

in the mitochondria, energy producing structures within 

cells, of B. procyonis was determined using polymerase 

chain reaction (PCR) methodologies (Xie and others, 

2011). Diagnostic PCR tests were also developed for 

Baylisascaris, and they can identify the parasite’s eggs in 

environmental samples or animal feces, as well as larvae 

recovered from paratenic host tissues or even in a single 

histopathologic section (Dangoudoubiyam and others, 

2009; Gatcombe and others, 2010; Jimenez Martinez and 

others, 2015). Standard PCR methods can distinguish B. 

procyonis from other parasites, including B. transfuga, but 

the tests do not distinguish B. procyonis from B. columnaris, which indicates how closely related these two species 

are (Kazacos, 2001; Dangoudoubiyam and others, 2009; 

Gatcombe and others, 2010). These two species can, however, be separated by DNA sequencing (Nadler and others, 

2010; Franssen and others, 2013), but these methods are not 

used routinely in assessing cases or outbreaks. Therefore, 

epidemiologic evidence of exposure to raccoons, skunks, 

or other animals and their feces remains very important 

for determining which Baylisascaris species is involved 

in a case or outbreak (Kazacos, 2001). Molecular methods 

are also being used in phylogenetic studies of the parasite 

(Nadler and Hudspeth, 2000; Nadler and others, 2010) and 

for genetic characterization and species identification of 

specimens recovered from raccoons (Blizzard, Davis and 

others, 2010; Blizzard, Yabsley and others, 2010) and other 

hosts (Tokiwa and others, 2014). As molecular methods 

become more widely available, they will increasingly be 

used to identify particular Baylisascaris species involved in 

clinical cases and outbreaks (Sato and others, 2004; Dangoudoubiyam and others, 2009).
Children and others may be 

exposed to raccoon fecal 

contamination at various 

sites in a yard. 

Dogs can also become 

infected and shed eggs 

in their feces.

Soil contamination.

Soil contamination.

Raccoon latrine on log.

Rodents and birds forage 

at latrines and become infected.

Latrine at 

base of tree.

Soil contamination at 

downspout.

Latrine 

on roof.

Feces and eggs may

wash to ground below.

Sandboxes and sandy 

play areas are possible 

sites of contamination.

Latrine on woodpile.

66 Baylisascaris Larva Migrans

Disease Ecology

The disease ecology of Baylisascaris larva migrans is 

straightforward, in that humans and animals become infected 

through accidentally ingesting infective eggs from the environment (Kazacos, 2001) (figs. 8 and 14). Thus, the ecology of 

the disease is directly related to the interaction of humans and 

animals with raccoons or other definitive hosts contaminating 

the environment, and in particular with their feces containing infective eggs. The longevity and resistance of infective 

eggs in the environment is the key feature in the transmission 

of baylisascariasis, because it ensures eventual contact and 

accidental ingestion of the eggs by animals or humans and the 

production of larva migrans.

In nature, the ability of B. procyonis to produce CNS disease in paratenic hosts has great survival value for the parasite, 

because altered behavior, debilitation, or death of paratenic 

hosts results in increased transmission of B. procyonis back 

to raccoons when they prey on infected smaller animals or 

scavenge infected carcasses (Tiner, 1953a,b; Kazacos and 

Boyce, 1989; Sheppard and Kazacos, 1997; Kazacos, 2001). 

Raccoons are opportunistic carnivores, and it is likely that 

Figure 14. Areas near homes where humans and animals are at risk of contacting raccoon feces containing B. procyonis eggs.

the increased pathogenicity of B. procyonis in paratenic hosts 

has been beneficial for the survival of B. procyonis over time, 

because of its importance in facilitating transmission and 

completion of the life cycle (Kazacos, 2001).

Because most B. procyonis eggs are found associated 

with raccoon latrines, contact and interaction of humans and 

animals with these sites is of considerable importance for 

transmission of the parasite (Page, 1998, 2013; Page and others, 1998, 1999, 2001a,b,c; Page, Anchor, and others, 2009; 

Kazacos, 2001; Smyser and others, 2010). In fact, raccoon 

latrines are central to the epidemiology and disease ecology of 

B. procyonis larva migrans, because they are the most dangerous areas for transmission of the parasite to humans and 

animals (Kazacos, 2001). Contact with other areas or articles 

contaminated with raccoon feces also poses a risk (fig. 14), as 

does contact with fecal material from dogs, skunks infected 

with B. columnaris, or other animals infected with Baylisascaris spp.. It is known that multiple raccoons will use a particular latrine and that individual raccoons will visit multiple 

latrines in their area (Hirsch and others, 2014). This ensures 

contamination of latrines with B. procyonis eggs and increases 

the risk of infection of paratenic hosts from latrine sites.
Disease Ecology 67

Latrines and Eggs—This is the Dangerous Part

Both animals and humans become infected with 

B. procyonis larvae by accidentally ingesting infective 

eggs from areas or articles contaminated with raccoon 

feces. These areas are often raccoon latrines, the preferred 

sites of raccoon defecation where large amounts of their 

feces and large numbers of B. procyonis eggs accumulate 

(Cooney, 1989; Page, 1998, 2013; Page and others, 1999; 

Page, Anchor, and others, 2009; Kazacos, 2001). Raccoon 

latrines can be identified by the accumulation of characteristic feces, which are dark and tubular, have a pungent odor, 

and often contain abundant seed material (Page and others, 

1998). They also contain abundant B. procyonis eggs, 

which present a real danger to animals and humans who 

may come into contact with them. Latrines in an area are 

used by multiple raccoons at different times, and individual 

raccoons will use multiple latrines (Hirsch and others, 

2014). Sampling latrines in an area for B. procyonis eggs 

provides important information concerning the potential 

risk of human and animal exposure to infection with this 

parasite (Kazacos, 2001; Page and others, 2005; Page, 

Anchor, and others, 2009).

In woodlots where raccoons are common, it is not 

unusual to find raccoon latrines and scattered feces on large 

logs and rocks, on stumps, on debris piles, at the base of 

trees, in the raised crotches and on large limbs of trees, 

and other horizontal structures (Yeager and Rennels, 1943; 

Stains, 1956; Cooney, 1989; Kazacos and Boyce, 1989; 

Page, 1998; Page and others, 1998) (A). Latrines can be 

found in various locations throughout the forest, particularly along streambeds and the banks of creeks and rivers 

(B). In one study of raccoons in a suburban park intercut 

by ravines, most latrines were found in the upper reaches 

of the park just inside the forest edge, near homes. These 

locations were presumably related to greater food resources 

and den sites nearby, compared to the densely wooded and 

steeper sloped areas of the park (Cooney, 1989). Other 

latrines were found along the streambed running through 

the park and near shelters and picnic areas.

Food sources related to humans are very important 

for supporting urban and suburban raccoon populations 

(Prange and others, 2003, 2004), and if raccoons are 

encouraged through feeding to frequent peoples’ yards and 

homes or have access to stored grain or animal feed, they 

often will establish latrines nearby (C) (Kazacos, 2001; 

Roussere and others, 2003; Page, Anchor, and others, 

2009). Latrines may be found in barn lofts, outbuildings, 

attics, garages, on woodpiles, decks, roofs, and other locations in the domestic environment (D) (Kazacos, 2001; 

Kazacos and others, 1983; Kazacos and Boyce, 1989; 

Rouserre and others, 2003; Page, Anchor, and others, 

2009). Raccoon latrines vary in size from small to quite 

large and extensive. Some of the largest latrines are in barn 

lofts, attics, and other choice locations of raccoon denning 

and defecation, where they may measure several yards 

across and 8–10 inches or more deep with feces (Cunningham and others, 1994; Kazacos, 2001). Interestingly, one 

study found a lower number of urban raccoons infected 

with B. procyonis as well as lower worm burdens in urban 

raccoons than rural raccoons, presumably due to urban 

raccoons’ greater dependence on human-related food rather 

than predation of paratenic hosts (Page and others, 2008). 

However, latrines containing abundant eggs were still common in urban and suburban landscapes, including peoples’ 

backyards (Roussere and others, 2003; Page, Anchor, and 

others, 2009).

Latrines can be surprisingly common in suburban 

communities where raccoons are common. In three communities in northern California, 244 raccoon latrines were 

found on 164 residential properties with a latrine density of 

3.5–8.8 per acre; nearly half (44–53 percent) of the latrines 

examined contained B. procyonis eggs, many of which 

were infective (Rouserre and others, 2003). In the Chicago 

suburbs, 61 of 119 backyards examined (51 percent) had 

1–6 latrines, and 23 percent of the latrines contained B. procyonis eggs (Page, Anchor, and others, 2009). Both studies 

indicate the potential risk of contact with raccoon latrines 

and B. procyonis eggs in domestic environments where raccoons are common. 

In areas of high prevalence, infected raccoons shed 

on average 20,000–26,000 B. procyonis eggs per gram of 

A. Raccoon latrines on a fallen 

log over a stream, on a split tree, 

at the base of a tree in a city 

park. (Photos by Kevin Kazacos 

and John Jacobson)
68 Baylisascaris Larva Migrans

Latrines and Eggs—This is the Dangerous Part

feces, and juvenile raccoons shed more eggs than adults 

(Kazacos, 1982, 1983a; Snyder and Fitzgerald, 1987). In 

an area of much lower prevalence, infected raccoons were 

shedding on average 17,527 eggs per gram of feces (Reed 

and others, 2012), which is still quite substantial. Many 

animals shed more than 100,000 eggs per gram of feces, 

and the highest reported egg shedding by a raccoon is 

256,700 eggs per gram of feces (Kazacos, 1983a). Thus, 

raccoon latrines and other contaminated areas can contain 

extraordinary numbers of infective eggs (E). At these concentrations, an animal or person would not have to ingest 

much material to take in a dangerous number of eggs that 

could cause severe clinical disease. Very young children 

who put soil and other materials into their mouths would 

be at the greatest risk of infection (Kazacos, 2000). Once 

the eggs reach infectivity, which takes a minimum of 11–14 

days (Sakla and others, 1989) but usually 2 or more weeks 

under natural conditions, they will remain viable for years, 

given adequate moisture from rainfall or other sources. 

On the other hand, eggs present in latrines in attics, barn 

lofts, and other hot, dry locations die from desiccation, but 

even under these conditions they will remain viable for 7 

months (Shafir and others, 2011). The eggs can also survive 

freezing, and they were viable after 8 weeks at –20 degrees 

Celsius (°C) (Garrison and Kazacos, 1994) and 6 months at 

–15 °C (Shafir and others, 2011).

Because large numbers of B. procyonis eggs are 

present at raccoon latrines, these areas become important long-term sources of infection. As first alluded to by 

Tiner (1952a), there is good evidence that paratenic hosts, 

particularly grain-eating rodents, become infected with 

B. procyonis by foraging for undigested seeds and other 

materials present in raccoon feces at latrines (Wirtz, 1982; 

Kazacos and Boyce, 1989; Sheppard and Kazacos, 1997; 

B. Large latrine in flood-debris laden tree, Deschutes River, Oregon. Closeup of same latrine showing extent of contamination 

with raccoon feces. (Photos by Kevin Kazacos)

Mammals and birds that visited raccoon latrines.

[Bold type indicates an active forager of seeds in raccoon feces. 

From Page (1998); Page and others (1999; 2001a)]

Mammals Birds

White-footed mouse Mourning dove.

Meadow jumping mouse White-breasted nuthatch.

Eastern chipmunk Brown thrasher.

Eastern fox squirrel Blue jay.

Eastern gray squirrel Hermit thrush.

American red squirrel Swainson’s thrush.

Southern flying squirrel Wood thrush.

Woodchuck Ovenbird.

Eastern cottontail Northern junco.

Virginia opossum Yellow-rumped warbler.

Raccoon Northern flicker.

Striped skunk American robin.

Long-tailed weasel Tufted titmouse.

Red fox Downy woodpecker.

Domestic cat House wren.

Domestic dog Black-capped chickadee.

White-tailed deer Cardinal.

Carolina wren.

Eastern screech owl.

Page, 1998; Page and others, 1999). Visitation to raccoon 

latrines by 17 species of mammals and 19 species of birds has 

been documented, and white-footed mice, eastern chipmunks, 

eastern fox squirrels, Virginia opossums, white-breasted 
Disease Ecology 69

C. Raccoon latrines around homes: at the base of a pine tree in a 

backyard in northern California; closeup of same latrine showing 

multiple ages of raccoon feces in latrine and feces with seeds; raccoon 

feces on a deck just outside the backdoor of a home in Indiana. (Photos 

by Kevin Kazacos)

D. Raccoon latrines on roofs and in or near barns: on a shed roof near a 

garage; on a tile roof of married student housing at a university in southern 

California; on straw bales in a barn in Indiana; on a 2- by-10-inch board 

under a corn crib associated with NLM in a child in upstate New York. 

(Photos by Kevin Kazacos and William J. Murray)

E. Infective B. procyonis eggs 

recovered from a raccoon 

latrine in Indiana. (Photo by 

Kevin Kazacos and Sam Royer)

Latrines and Eggs—This is the Dangerous Part

nuthatches, and hermit thrushes were documented actively 

foraging for undigested seeds (Page, 1998; Page and others, 

1999, 2001a). Visitation by white-footed mice was significantly greater when corn, their most highly preferred seed 

type, was present in raccoon feces. Caching of raccoon feces 

by white-footed mice (Page, 1998) and Allegheny woodrats 

(McGowan, 1993; LoGiudice, 2001; Page and others, 2012; 

K.R. Kazacos and others, unpub. data, 1995) has also been 

documented, and such behavior has been linked to extirpation of Allegheny woodrats from parts of their northeastern 

range where raccoon populations have increased (McGowan, 

1993; LoGiudice, 2003) (See “Plight of the Allegheny Woodrat”). Animals could also become infected while investigating a latrine site, or indirectly through grooming, after having 

become contaminated at a latrine (Sheppard and Kazacos, 

1997; Page and others, 1999).

Humans, primarily young children, have become 

infected and developed fatal or severe CNS disease after 

ingesting material from or near raccoon latrines. This has 

taken place in the domestic environment in and around 

homes, on farms, in county parks or other areas along rivers, 

in childcare facilities, and group homes for developmentally 

disabled individuals (Huff and others, 1984; Fox and others, 

1985; Cunningham and others, 1994; Park and others, 2000; 

Rowley and others, 2000; Moertel and others, 2001; Kazacos, Gavin, and others, 2002; Gavin and others, 2002; Chun 

and others, 2009; Hajek and others, 2009). Several cases 

in children have involved contact with raccoon latrines in 

sandboxes or sandy play areas at their homes, at the base of 

trees in yards, or other locations with easy access by young 

children. Pica and geophagia by young children increase 

the likelihood of infection from such areas and have been 

incriminated in a number of these cases. It is important 

to realize that not only do curious young children find the 

material associated with raccoon latrines to be fascinating, but children younger than a certain age do not have an 

aversion to ingesting that material, resulting in potentially 

life-threatening infection with B. procyonis (Kazacos, 2001, 

2002; Gavin and others, 2002; Murray and Kazacos, 2004; 

Sanders, 2015).
70 Baylisascaris Larva Migrans

In addition to ingestion of infective eggs, humans or 

animals may also become infected with Baylisascaris larvae through ingestion of larvae in paratenic hosts. Paratenic 

transfer of larvae by ingestion of host tissues is well known for 

Toxocara and some other carnivore ascarids (Akao and Ohta, 

2007; Hoffmeister and others, 2007; Yoshikawa and others, 

2008), but it has not been examined for Baylisascaris. Nonetheless, it is a possible means of infection with larvae of these 

parasites, albeit it would be less important overall than ingestion of infective eggs. Therefore, any paratenic hosts destined 

for human consumption, especially wild mammals and birds, 

should be cooked thoroughly to kill any Baylisascaris larvae 

and other infectious agents that might be present. Another 

way animals, and probably humans, may become infected 

with Baylisascaris LM is through transplacental transfer of 

larvae undergoing somatic migration in pregnant females, as 

occurred in newborn lambs in Idaho (Anderson, 1999). Similar to the situation for Toxocara infection in dogs (Kazacos, 

1991), Baylisascaris larvae apparently are stimulated and (or) 

activated by the hormones of pregnancy.

People and animals are at risk of infection with B. procyonis in essentially all geographic areas where raccoons are 

found (Kazacos, 2001). Except for a few localized or regional 

areas where raccoons may not be infected with the parasite, 

B. procyonis is generally found to some extent in raccoons 

wherever they occur (table 5). In many urban and suburban 

areas, raccoon populations are large because of abundant food 

sources intentionally or unintentionally provided by humans, 

numerous den sites, and a lack of natural predators (Hoffman and Gottschang, 1977; Kidder, 1990; Rosatte and others, 

1991; Riley and others, 1998; Smith and Engeman, 2002; 

Gehrt, 2003; Prange and others, 2003, 2004; Page, 2013). Raccoons in such areas have higher reproductive rates and, with 

abundant available food, maintain strong, viable populations; 

unfortunately, this leads directly to increased environmental 

contamination with raccoon feces and B. procyonis eggs in 

these areas. In urban and suburban areas, raccoons are commonly considered a nuisance (de Almeida, 1987; Bluett and 

others, 2003) and frequently cause property damage and forage on refuse in garbage cans and dumpsters in neighborhoods 

and parks (Prange and others, 2004). Raccoons also scavenge 

food left out for dogs and cats (Roussere and others, 2003). It 

is not uncommon to see raccoon latrines established throughout such environments, posing a risk of Baylisascaris LM to 

people and animals living there (Roussere and others, 2003; 

Page and others, 2009a; Page, 2013).

Wild raccoons are also a well-recognized nuisance in and 

around zoos and other animal facilities, where they are responsible for considerable property damage and widespread fecal 

contamination. Raccoons are excellent climbers and will readily establish latrines in elevated locations, posing unique problems for zoos using large wire “roundhouse” enclosures and 

for people who may not notice rooftop contamination. Raccoon latrines in open exhibits or on the tops of roundhouse or 

other enclosures, as well as contaminated logs and tree limbs 

placed into exhibits, have resulted in numerous infections with 

B. procyonis (Armstrong and others, 1987, 1989; Kazacos 

and Boyce, 1989; Stringfield and Sedgwick, 1997; Kazacos, 

2001; C.L. Eng and K.R. Kazacos, unpub. data, 1997) (tables 

10–20). Latrines on rooftops are common (Roussere and others, 2003), can be quite extensive, and pose a threat because 

rain runoff will carry fecal material and eggs through rain 

gutters or directly to ground areas below, where they may 

be contacted by people or animals. This scenario led to the 

exposure of over 80 children in a childcare facility in southern 

California, where an infant boy contracted severe NLM due to 

B. procyonis; several other children were seropositive, indicating infection (Schultz, 2002; Murray and Kazacos, 2004; K.R. 

Kazacos and others, unpub. data, 2002). Similarly, DeBrazza’s 

monkeys contracted NLM in Indiana from raccoon feces and 

eggs raining down into their living area from a latrine on the 

top of their wire roundhouse enclosure (C.L. Eng and K.R. 

Kazacos, unpub. data, 1997).

Ample opportunities exist for contact and infection of 

domestic animals with B. procyonis. Fatal CNS disease due to 

B. procyonis has been seen in a wide variety of domestic animals, including pet dogs, rabbits, North American porcupines, 

canaries, parrots, and farm-raised rabbits, chinchillas, foxes, 

poultry, quail, pheasants, and large flightless birds (Kazacos, 

2001) (tables 10–20). Infection with B. procyonis has been 

linked many times to the use of straw, hay, and feed contaminated by wild raccoons denning and (or) defecating in such 

materials (Richardson and others, 1980; Kazacos, Kazacos, 

and others, 1982; Kazacos, Reed, and others, 1983; Kazacos, 

Reed, and Thacker, 1986; Armstrong and others, 1987, 1989; 

Sanford, 1991; Van Andel and others, 1995; Campbell and 

others, 1997; Pessier and others, 1997; Lennox and others, 

2015). In agricultural areas, it is common to find raccoon 

latrines in barns and other outbuildings or on old farm machinery, especially if stored grain or other animal feed is present 

nearby. Other cases have been linked to direct contamination 

of animal facilities and enclosures by raccoons and to the use 

of contaminated cages and enclosures that previously kept or 

housed raccoons (Schueler, 1973; Church and others, 1975; 

Koch and Rapp, 1981; Reed and others, 1981; Larson and 

Greve, 1983; Myers and others, 1983; Dixon and others, 1988; 

Medway and others, 1989; Fitzgerald and others, 1991; Coates 

and others, 1995; Garlick and others, 1996; M.A. Nieves and 

others, unpub. data, 1989). Other infections have occurred 

in wildlife rehabilitation and animal dealer facilities following the use of contaminated cages or enclosures, hay, or feed 

(Pigage and others, 1983; Dixon and others, 1988; K.R. Kazacos, unpub. data, 1983, 1984). A large outbreak occurred in 

guinea pigs in a “closed” American Association of Laboratory 

Animal Care-accredited research animal facility, due to the 

unwitting introduction of contaminated bedding that had been 

stored in an outbuilding accessed by raccoons (Van Andel 

and others, 1995). In the United States, Rhesus macaques at a 

research animal breeding facility became infected, presumably 

due to direct contamination of large enclosures by wild raccoons (Gozalo and others, 2008), and in Japan, cases in Japanese macaques were linked to contamination from raccoons 
Disease Ecology 71

also kept in a safari-style zoo (Sato and others, 2005). Zoo 

contamination and infection is common, as evidenced by 29 

percent of nonhuman primates at six major zoos in the United 

States testing positive for antibodies against Baylisascaris

during blood serum tests (Zimmerman and others, 2010). An 

extensive outbreak involving B. columnaris infection in three 

species of marmosets and tamarins at a zoo was linked to two 

infected skunks kept in the same exhibit, presumably to make 

it more “natural” (Huntress and Spraker, 1985; K.R. Kazacos 

and P.L. Wolff, unpub. data, 1986). Marmosets and tamarins 

spend much of their time on the ground foraging, which apparently led to infection with Baylisascaris (Pessier and others, 

1997). 

Animals may die rapidly or over longer periods of time 

due to Baylisascaris NLM. In the largest outbreak of NLM 

ever recorded, 640 bobwhite quail died following exposure to 

pens and straw contaminated by wild raccoons (K.R. Kazacos 

and others, unpub. data, 2002). This rivaled an earlier outbreak 

in which 622 chickens in a poultry facility died over a 7-week 

period, following the use of straw litter contaminated by raccoon feces (Richardson and others, 1980). In another incident, 

85 bobwhite quail were placed in a 12-by-24 foot dirt pen that 

had housed 3 young pet raccoons and was heavily contaminated with Baylisascaris eggs; all of the quail died (Reed and 

others, 1981; Kazacos, 1982). In a fourth dramatic outbreak, 

all 10 pet birds in a mixed collection of parrots and conures 

succumbed quickly to massive B. procyonis infection acquired 

from contaminated feed. Feral raccoons had contaminated a 

seed mixture stored in a large container in the owner’s garage, 

and this material was unwittingly fed to the birds, resulting 

in severe disease (Lennox and others, 2015). Other outbreaks 

have been insidious, with sporadic or constant low-level losses 

of animals from NLM. This type of outbreak took place on 

a pheasant ranch in Wisconsin, where 1–2 percent of young 

birds were affected over a 2-year period (Kazacos, Reed, and 

Thacker, 1986). The young birds were raised in groups in a 

barn, whereas older pheasants kept in outside wire pens were 

not affected. Problems began when the bedding for the young 

birds was changed to straw, which was obtained from a neighbor’s barn and subsequently found to be contaminated with 

raccoon feces and B. procyonis eggs. Subtle disease and losses 

have been seen in numerous other instances in animals with 

isolated or low-level exposures and indicate the commonality 

of low-level, covert infection in both animals and humans.

To date, 25 cases of Baylisascaris NLM have been 

reported in humans (table 8) (Kazacos, 2001; Kazacos, Gavin, 

and others, 2002; Schultz, 2002; Murray and Kazacos, 2004; 

Cheney, 2005; Chris, 2005; Gavin and others, 2005; Pai and 

others, 2007; Reilly, 2008; Chun and others, 2009; Hajek and 

others, 2009; Moore, 2009; Kelly and others, 2009, 2012; 

Mehta and others, 2010; Perlman and others, 2010; Ciarlini 

and others, 2011; Hung and others, 2012; Haider and others, 

2012), and at least a dozen additional cases are known (C.J. 

Crosley and others, unpub. data, 2005; Crosley and Kazacos, 

2005; K.R. Kazacos, unpub. data, 1995–2010). Six of these 

cases resulted in death from overwhelming infection. Most 

cases have involved infants.

The first two recognized cases of B. procyonis NLM in 

humans were fatalities in infants who became infected within 

their homes, not in the outside environment. The first case was 

a 10-month-old boy in southeastern Pennsylvania who rapidly 

deteriorated into a comatose state, dying 14 months later. He 

was believed to have ingested raccoon feces from open fireplaces within the home, contaminated by raccoons denning in 

the chimneys (Huff and others, 1984). The other case was an 

18-month-old boy with Down syndrome in northeastern Illinois who developed severe disease that progressed quickly to 

death; infection resulted from chewing on pieces of bark from 

contaminated firewood brought into the home. The bark came 

from logs originating as downed timber in a local woodlot that 

had many established raccoon latrines (Fox and others, 1985; 

Sanders, 2015). Another early case was a 13-month-old boy in 

upstate New York who became infected on a dairy farm while 

playing in the soil near a large raccoon latrine under a corn 

crib (Cunningham and others, 1994). In a case in northern California, many infective B. procyonis eggs were recovered from 

the sand in a child’s swing set play area and from 21 raccoon 

latrines found on the property and an adjacent vacant lot (Park 

and others, 2000). In the Chicago suburbs, two infant boys 

became infected in their own or neighbor’s yards (Gavin and 

others, 2002). Both children regularly put fingers and objects, 

including backyard dirt, into their mouths. One boy became 

infected while playing “fort” in a cluster of small trees that 

contained an active raccoon latrine, from which he ate material. An 11-month-old child in southern California became 

infected in a university childcare facility, where feces and eggs 

from large raccoon latrines on the roof of the building washed 

off into the infant play area below; this child also mouthed 

objects and ate dirt (K.R. Kazacos, unpub. data, 2002; 

Schultz, 2002; Murray and Kazacos, 2004; Sanders, 2015). 

A more recent case in an infant girl in Missouri (Mehta and 

others, 2010) involved probable ingestion of contaminated soil 

and (or) raccoon feces in a picnic area in a small rural county 

park along a river, where there was abundant raccoon activity 

and contamination (K.R. Kazacos, unpub. data, 2007). Due to 

their tendency for oral sampling of their environment, including pica and geophagia, infants are at markedly increased risk 

of severe infection (Kazacos, 2000, 2001; Murray and Kazacos, 2004; Gavin and others, 2005; Kazacos and others, 2013; 

Singaravelu and others, 2016).

Severe CNS disease developed in a 63-year-old man 

in northern California after he worked under his house in a 

raccoon feces-contaminated area and ate lunch there without 

handwashing (C. Langelier and others, written commun., 

2015). At least six cases have occurred in developmentally 

disabled, autistic, mentally retarded, or demented adolescents 

or adults, and a seventh in a teenage drug abuser, all of whom 

exhibited altered behavior (pica, geophagia) that put them at a 

greater risk of infection (Fox and others, 1985; Cunningham 

and others, 1994; Moertel and others, 2001; Kazacos, Gavin, 
72 Baylisascaris Larva Migrans

and others, 2002; Chris, 2005; Chun and others, 2009; Hajek 

and others, 2009; Ciarlini and others, 2011; Hung and others, 

2012). In one such case in southern California, a 17-year-old 

in a group home for developmentally disabled adolescents 

became comatose and died following ingestion of large 

amounts of sand from a contaminated sandbox at the facility; a 

brain biopsy was positive for larvae (Kazacos, Gavin, and others, 2002). In Oregon, a teenager somehow became infected 

during or following drug abuse while camping along a river in 

a remote area (Chun and others, 2009).

Although most cases of NLM and OLM–DUSN are 

related to contact with contaminated areas in the outside 

environment, infected animals kept as pets or in zoos and 

other collections may also be a source of infection. Cases of 

clinical OLM–DUSN were seen in a teenager in Kentucky and 

an adult in Germany several weeks after they acquired pet raccoons (Raymond and others, 1978; Kuchle and others, 1993). 

Contact with areas or articles contaminated by infected pet 

skunks, other procyonids, bears, or pet dogs could also result 

in infection (Kazacos, 2001, 2006; Kazacos, Kilbane, and others, 2011). Skunks are raised for the exotic pet trade on large 

breeding farms in Iowa and elsewhere, and young descented 

skunks are sold in late spring in pet stores in states where 

it is legal. Small breeders also contribute to the pet trade in 

these animals, which may be sold infected with B. columnaris

(K.R. Kazacos, unpub. data, 1997–99). Kinkajous and coatis 

are imported from South America and (or) raised in breeding 

facilities in Florida and elsewhere and then sold throughout 

the United States and in other countries in pet stores or by private owners or small breeders; these animals may be infected 

with B. potosis or other species (Kazacos, Kilbane, and others, 

2011; Taira and others, 2013; Tokiwa and others, 2014; Parkanzsky, 2015). Raccoons are also bred and raised by animal 

dealers who sell the young as exotic pets and export them to 

other countries for pets or placement in zoos; thousands of raccoons have been introduced into Japan in this way (Miyashita, 

1993) and hundreds into China (Xie and others, 2014). Other 

hosts of Baylisascaris species, including other procyonids 

such as ringtails and olingos, or bears, bred and sold as pets or 

for collections are also of concern. All of these animals pose a 

potential threat for introducing Baylisascaris into households, 

animal facilities, and local environments and exposing people 

and animals to possible infection.

Points to Ponder

Baylisascariasis is an excellent example of a disease 

where human and domestic animal infection results from close 

association with a common wildlife species, the raccoon, 

which many people consider to be both engaging and relatively harmless. It is a disease to be reckoned with in humans 

and animals alike, as individuals, groups, as well as populations. Cases may involve only one or a few affected individuals or, in heavier exposures, entire groups may be affected 

with mortality at or near 100 percent (Richardson and others, 

1980; Reed and others, 1981; Kazacos, Reed, and others, 

1983; A.M. Lennox and others, unpub. data, 1996; Lennox and 

others, 2015). The remarkable pathogenicity of B. procyonis

and its ability to infect a wide variety of hosts are such that 

once it becomes established in an area, the consequences of its 

presence become known through clinical cases of neurologic 

and ocular disease affecting a wide range of animals as well as 

people (Kazacos, 2001; Page, 2013). Depending on the prevalence of infection in raccoons and the animal species affected, 

B. procyonis could have devastating effects on particular 

species or populations, even at relatively low environmental 

levels. Thus, it is very important for wildlife biologists and 

public health and medical personnel to be aware of the parasite 

and what it can do clinically and to urge caution and restraint 

pertaining to the translocation of raccoons to new areas for 

whatever purpose. Because current high raccoon densities in 

different areas maintain B. procyonis at high levels, indiscriminate translocation of raccoons by hunting clubs, pet owners, 

and others could introduce the parasite into new areas, where 

it could become established and pose a threat to indigenous 

birds and mammals, including humans (Kazacos and Boyce, 

1989; Kazacos, 2001; Page, 2013).

Animals other than raccoons, including other procyonids and domestic dogs that are being bred and sold in North 

America and elsewhere as pets, could also be involved in 

introducing and (or) spreading B. procyonis and related parasites to new areas. In addition to kinkajous and coatis infected 

with Baylisascaris spp., several dozen dogs have also been 

found to have patent infections with B. procyonis in areas 

where it occurs in raccoons and, due to the commonality of pet 

travel, infected dogs could spread B. procyonis to new areas 

(Kazacos, 2001, 2006). These situations point to the need for 

increased veterinary vigilance, routine deworming of animals, 

and better regulations for animal health certifications and 

prophylactic deworming of potential definitive hosts before 

importation, exportation, or interstate transport. Infected 

paratenic hosts carrying larvae could also introduce and spread 

B. procyonis if they are translocated from enzootic areas and 

released, provided that competent definitive hosts are present 

in the new location.

Early field studies estimated that B. procyonis was 

responsible for as much as 5 percent of rodent mortality in 

Illinois woodlots where raccoons were common (Tiner, 1954). 

More recently, larva migrans due to Baylisascaris was found 

to be surprisingly common in white-footed mice collected in 

woodlots in agricultural north-central Indiana, where 28–29 

percent of mice were infected (Page and others, 2001c; Beasley and others, 2013). Similarly, 33 percent of mice in urban 

forest preserves in Chicago, Illinois were infected with B. procyonis (Kellner and others, 2012). Based on the pathogenicity 

of B. procyonis larvae in mice, levels of clinical disease similar to or greater than those seen by Tiner (1954) could easily 

be expected. Landscape type as well as land-use are important 

attributes affecting prevalence of B. procyonis, as they affect 

host density, the behaviors and interactions of raccoons and 

paratenic hosts, the availability of food resources, contact 
Disease Prevention and Control 73

rates and interactions of paratenic hosts with latrine sites, and 

other factors (Page, 2013). In Indiana, raccoons in agriculturedominated landscapes had a higher prevalence of B. procyonis

infection than those in forest-dominated landscapes (Page and 

others, 2001c). Similar results were seen in Wisconsin, where 

infected raccoons had more agricultural and fewer forested 

areas in their home ranges than did uninfected raccoons (Samson and others, 2012). This type of habitat correlates with a 

greater abundance of food resources, including paratenic hosts 

such as white-footed mice (Nupp and Swihart, 2000; Page and 

others, 2010c), and favors transmission to paratenic hosts in 

these areas. Despite their sometimes lower prevalence of B. 

procyonis infection, urban raccoons remain important contributors to environmental contamination and thus transmission of 

the parasite because of their high numbers (Page and others, 

2008; Kellner and others, 2012; Page, 2013). 

It is estimated that hundreds, if not thousands, of clinical 

cases presently occur each year in animals in North America, 

especially cases affecting granivorous rodents and birds. The 

vast majority of clinically affected wild animals would escape 

human notice because of their small size, isolated location, 

and the fact that they would be more readily taken by predators and scavengers. Numerous wildlife species ranging from 

songbirds to large mammals forage at raccoon latrines and 

are thus in a direct line to become infected with this parasite 

(Page and others, 1999, 2001a,b). NLM due to B. procyonis

was a common finding in wild animals in southern California 

(Evans, 2002), where the prevalence of the parasite in raccoons was also high (Evans, 2001).

The potential impact of Baylisascaris infection on natural 

populations of wildlife, including threatened and endangered 

species, is well demonstrated by the plight of the Allegheny 

woodrat in the northeastern United States. The demise of 

the woodrat in this region serves as an excellent example of 

the potential effects of B. procyonis on animal populations, 

following the introduction and (or) increase in raccoons in an 

area (Kazacos, 2001; Page, 2013). It is also a glaring example 

of the consequences for humans and domestic animals of living in close association with infected raccoons. People in such 

areas need to be made aware of parasites such as B. procyonis, 

so that they can take appropriate precautions concerning wildlife, the outside environment, and especially raccoon latrines, 

and follow basic preventive measures to protect themselves, 

their families, and their animals from infection. 

Disease Prevention and Control

Prevention of B. procyonis infection in humans and 

animals depends on preventing ingestion of infective eggs 

from environmental sources contaminated by raccoons. 

This, in turn, requires preventing or limiting raccoon fecal 

contamination and dealing with it effectively once it has 

occurred. Because of the seriousness of Baylisascaris infection in humans and animals, and the present lack of effective 

treatment for NLM (see “Treatment,” below), prevention of 

infection with Baylisascaris is of utmost importance (Kazacos, 2001). Three key elements for preventing and controlling 

Baylisascaris infections in humans and animals include (1) 

reducing environmental contamination with infective eggs; 

(2) preventing contact with contaminated areas or articles; and 

(3) educating people about Baylisascaris as a cause of human 

and animal disease so that they can take appropriate precautions (Kazacos, 1991, 2000, 2001; Kazacos and Boyce, 1989). 

These approaches should be carried out together as part of 

an overall prevention and control program and should apply 

not only to raccoons, but as well to other procyonids, dogs, 

skunks, and other definitive hosts of Baylisascaris spp.

Reducing Environmental Contamination

Environmental contamination with Baylisascaris eggs 

in an area is reduced by treating, removing, and (or) relocating infected raccoons and skunks (LoGiudice, 1995; Kazacos, 

2001; Page and others, 2011). Keeping raccoons and skunks as 

pets should be strongly discouraged, particularly in households 

with young children (Kazacos, 2000, 2001). Precautions are 

also important when keeping kinkajous, coatis, ringtails, other 

procyonids, or bears as exotic pets. Dogs infected with B. procyonis can also contaminate peridomestic areas. Anthelmintic 

treatment of raccoons, other procyonids, skunks, bears, and 

dogs kept as pets or for other reasons is easily accomplished, 

but it must be done adequately in order to prevent contamination with eggs (Kazacos, 2001).

Depopulation and Removal

Wild raccoons are involved in most Baylisascaris infections (Kazacos, 2001). The most straightforward method of 

dealing with infected raccoons in an area is through depopulation and removal (Kazacos, 2001). This immediately reduces 

new environmental contamination in an area and is best 

combined with latrine cleanup and decontamination. Depending on a particular state’s regulations as to how these animals 

are dealt with, trapped animals may be relocated to distant 

sites or euthanized. It is not uncommon for suburban zoos to 

have ongoing wildlife control programs in an effort to reduce 

and control nuisance wildlife and the diseases they carry. Trapping and removal may be applied on a localized basis, directed 

against particular nuisance animals, or on a broader scale.

Typically, these efforts must be sustained or periodically 

applied, because removal of animals creates niche and population voids, which will be filled by new raccoons moving into 

the area. Consider the situation in the late 1990s, on the Monterey Peninsula in northern California, where the adjacent cities of Pacific Grove and Carmel had ongoing nuisance raccoon 

problems, with frequent complaints from citizens (Roussere 

and others, 2003). A case of severe NLM in a child in Pacific 

Grove in August 1998 (Park and others, 2000) prompted a 

program of trapping and euthanasia of nuisance raccoons, as 
74 Baylisascaris Larva Migrans

Plight of the Allegheny Woodrat

The Allegheny woodrat (A) is threatened or endangered 

through much of its Northeastern and Midwestern range. It 

has become extirpated from New York, Connecticut, and 

much of New Jersey and continues to decline in other areas 

(Balcom and Yahner, 1996; LoGiudice, 2006). Although a 

number of factors may be involved, declines in woodrats in 

the Hudson Highlands region of New York were convincingly linked to neural larva migrans (NLM) caused by B. 

procyonis, related to increases in the raccoon population 

in the area over time (Kazacos, 2001; LoGiudice, 2003). 

Similar associations were also made with raccoon fecal 

contamination in woodrat habitat in southern Indiana (Page 

and others, 2012).

Based on long-term naturalist’s records at the Mohonk 

Preserve in New Paltz, N.Y., raccoons were considered a 

rarity there in 1923–32, but they were abundant by the end 

of 1949 and subsequently (Smiley, 1977a). Declines in the 

woodrat population had also been noted, and it was thought 

that this might have been related somehow to the increase 

in raccoon numbers (Smiley, 1977b). In 1986, abnormal 

behavior was documented in various animals in the preserve, 

including gray squirrels and deer mice found circling or 

unable to climb trees (Smiley and Huth, 1986). B. procyonis

NLM was suggested as the probable cause of this abnormal 

behavior, in addition to its possible role in the extirpation of 

woodrats from the preserve between 1959 and 1977 (W.B. 

Stone, written commun., 1998; Kazacos, 2001). Woodrats 

became extirpated from the state in 1987, and subsequent 

woodrat release studies undertaken by the New York Department of Environmental Conservation at Mohonk in 1991 

indicated that B. procyonis was the likely cause of the loss of 

woodrats from the region, in combination with other factors 

(McGowan, 1993). 

The problem stemmed from both habitat type and the 

inherent behavior of woodrats. Allegheny woodrats prefer 

steep rock and boulder talus slopes, escarpments, cliff ledges 

and caves, with deep secluded areas for nesting (B, C). In 

eastern New York State, these areas are also very attractive 

to raccoons, which used them as den and latrine sites (D) 

(Kazacos, 2001). As the raccoon population increased, so 

did raccoon latrines, with increased chances of woodrats 

contacting those latrines and B. procyonis eggs. Uninfected 

woodrats from West Virginia released into these areas, as 

well as their offspring, died of Baylisascaris NLM, evidence 

of which was documented in all 11 woodrats recovered 

and examined; 4 of these animals had exhibited abnormal behavior when they were live-trapped (McGowan, 

1993). Baylisascaris NLM was also identified in Allegheny 

woodrats in southern Indiana (E) (K.R. Kazacos and S.A. 

Johnson, unpub. data, 1996) and south-central Pennsylvania 

(J. Wright and others, unpub. data, 1998), and in woodrats 

released in New Jersey (LoGiudice, 2003) and southern 

Indiana (Smyser, Johnson, and others, 2013). Studies of 

raccoon latrine associations in woodrat habitat in southern 

Indiana found a correlation between positive latrines containing infective eggs and lower woodrat numbers (Page and 

others, 2012).

Woodrats, a type of packrat, are well known for their 

caching behavior, and in New York, New Jersey, and Indiana, the extensive caching of raccoon feces by woodrats was 

documented, indicating a direct behavioral link to raccoon 

latrines and B. procyonis infection. Caching behavior was 

examined further and revealed that whereas a white-footed 

mouse tends to pick apart feces and take out seeds immediately upon finding fresh raccoon feces, a woodrat prefers 

older dried feces (mean, 21 days old) and would carry the 

A. Allegheny woodrat in a 

cave in southern Indiana 

along the Ohio River. (Photo 

by Kevin Kazacos)

B. Talus boulder slope habitat 

of Allegheny woodrat in eastern 

New York State. (Photo by 

Kevin Kazacos)

C. Cliff ledge 

habitat of 

Allegheny 

woodrat in 

southern 

Indiana. 

(Photo by 

Kevin Kazacos)

D. Raccoon latrine in talus 

boulder habitat in eastern 

New York State. (Photo by 

Kevin Kazacos)

E. Allegheny woodrat from southern 

Indiana with Baylisascaris NLM, 

showing severe head and body 

tilt, extension of the forelimbs, 

and falling over. (Photo by Scott A. 

Johnson)
G. Allegheny woodrat with NLM following experimental infection with 

B. procyonis. Photos show arching of the head and neck, extension and 

rigidity of the forelimbs, and paddling movements while lying down. 

(Photos by Kevin Kazacos and Sam Royer)

F. Portion of raccoon feces 

cached by a woodrat in its 

midden in southern Indiana. 

(Photo by Kevin Kazacos and 

Sam Royer)

Disease Prevention and Control 75

Plight of the Allegheny Woodrat

entire piece of feces back to its den and cache it in its midden (F) (LoGiudice, 2001). Carrying raccoon feces in their 

mouths, as well as later consumption of seeds present in 

older fecal material, puts woodrats at great risk of infection 

with B. procyonis eggs, which would have reached infectivity by then.

Experimental studies on the susceptibility of woodrats to B. procyonis found them to be highly susceptible to 

NLM (G) (K.R. Kazacos, unpub. data, 1997). Because one 

larva in the brain of a susceptible small rodent is usually 

fatal (Tiner, 1953a,b; Sheppard and Kazacos, 1997), and a 

rodent may die following infection with a surprisingly low 

number of eggs (Dubey, 1982), woodrats are at particular 

risk when they are repeatedly exposed to even low numbers 

of eggs. Thus, they would be at risk of NLM even in areas 

of low B. procyonis prevalence in local raccoons and latrine 

sites (Page and others, 2012). Studies of woodrats released 

in New Jersey showed that those in less contaminated sites 

were at risk of NLM, but that the hazard of death increased 

2.67 times for those in highly contaminated sites (LoGiudice, 2003). Every additional raccoon latrine found per 1 

hour of search time increased the death hazard to woodrats 

by 10 percent (LoGiudice, 2003).

The unique plight of the Allegheny woodrat demonstrates that Baylisascaris can have a significant impact 

on certain indigenous wildlife, especially species that are 

already compromised due to low population numbers and 

(or) habitat loss. At particular risk are grain-eating species 

that may forage at raccoon latrines (Page and others, 1999, 

2001a), especially those like the woodrat that normally 

cache materials.

Therefore, it becomes very important to consider the 

consequences of raccoon translocation into new areas, as 

well as their natural dispersal and population increases, 

because all could affect B. procyonis prevalence and its 

effect on other species (Page, 2013). If other species are 

threatened or endangered like the Allegheny woodrat, the 

consequences may be dire (Kazacos, 2001; LoGiudice, 

2003; Page, 2013). Prevention or limitation of raccoon 

translocations as well as thorough deworming of translocated raccoons would be important in reducing the spread 

of B. procyonis to new areas and its consequences for other 

species. Other interventions being studied to prevent the 

impact of B. procyonis on woodrat populations (as well 

as other species, including humans) include the use of 

anthelmintic-laced baits to deworm local raccoon populations (Kazacos, 2001; Page, 2013). Limited experiments in 

New Jersey indicated that this might work, because following the use of deworming medications in baits, B. procyonis

worms and tracking dye were found in raccoon feces at 

latrines (LoGiudice, 1995). A study in Indiana showed that 

bait deworming decreased the prevalence of B. procyonis eggs in raccoon latrines as well as larval infections in 

white-footed mice in areas where the raccoons were treated 

(Page and others, 2011). Another study found that the use 

of anthelmintic-containing baits increased the success of 

Allegheny woodrat translocations into previously occupied 

sites (Smyser, Johnson, and others, 2013). One problem 

not usually considered with this approach, however, is that 

the eggs in worms passed by treated raccoons will still 

develop to infectivity and be present in latrines, available for 

reinfection of raccoons as well as infection of other animals, 

including woodrats. However, bait deworming is a step in 

the right direction, and regular treatment combined with 

regular latrine cleanup and decontamination in an area may 

have a measurable impact on decreasing the extent of this 

problem, particularly in smaller targeted areas including in 

and around known woodrat habitats.
76 Baylisascaris Larva Migrans

well as educational efforts aimed at discouraging people from 

feeding or providing shelter for these animals. These efforts, 

which were instituted in Pacific Grove but not in Carmel, 

prompted lawsuits from animal rights groups, cease-and-desist 

orders, and finally legal resumption of trapping when, for public health reasons, opposition was denied in local courts (W.J. 

Murray, pers. commun., 2001). Subsequent studies found a 

marked difference in latrine densities in the two cities, with 

8.7 latrines per hectare in Pacific Grove versus 21.7 per hectare in Carmel, apparently related to their different respective 

efforts to control local raccoons (Roussere and others, 2003).

Anthelmintic Baiting

Other than trapping and removal, B. procyonis prevalence 

in an area may be reduced by using baits containing anthelmintics to deworm wild raccoons, similar to the bait treatment 

of Echinococcus in foxes in Europe or the use of baits for 

rabies vaccination of wildlife (Kazacos, 2001). Although the 

frequency, timing, and logistics of such treatments related to 

their overall effectiveness are not well known, encouraging 

results were seen in studies done in woodrat habitat in central 

New Jersey (LoGiudice, 1995), in agricultural areas of northcentral Indiana (Page and others, 2011), and in woodrat habitat 

in southern Indiana (Smyser, Johnson, and others, 2013; 

Smyser, Page, and others, 2013). In New Jersey, raccoons 

were baited with piperazine at two sites. The number of scats 

containing eggs was significantly reduced overall at the treatment sites compared to the control sites, but it was impossible 

to identify the contributions of individual animals to feces at 

latrines. However, successful treatment was indicated by finding adult Baylisascaris in two of three scats that also contained 

a fluorescent marker indicating bait consumption. Nothing 

could be concluded as to the effects of anthelmintic baiting on 

overall prevalence of the parasite, because too few raccoons 

were trapped and examined for eggs. 

These results of anthelmintic treatment of raccoons in 

woodrat habitats were strongly supported by the studies in 

Indiana (Page and others, 2011; Smyser, Johnson, and others, 2013; Smyser, Page, and others, 2013). In the first study, 

raccoons were baited monthly with pyrantel pamoate, a 

deworming agent used to treat roundworm infections, after 

first removing and torch-sterilizing latrines in eight forest 

treatment patches. The prevalence of B. procyonis eggs in 

latrines in treatment patches declined by over threefold in all 

sampling periods after baiting, and during one year, there was 

a significant decline in the prevalence of B. procyonis larvae 

in white-footed mice in treatment areas compared with control 

patches. This was the first study to show a direct effect on 

larva migrans in paratenic hosts in an area following the bait 

deworming of raccoons, and served as a baseline for later 

studies assessing the efficacy of baiting for sustained control 

of Baylisascaris in wild raccoons.

In the other studies, monthly baiting with pyrantel 

pamoate was conducted in nine sites in woodrat habitat 

(limestone and sandstone cliffs) in southern Indiana that 

included all five known extant woodrat populations in the 

state. Anthelmintic baiting for 24 months led to reductions in 

prevalence of B. procyonis eggs in latrines from prevalences 

of 9–11 percent before treatment to near 0 percent after treatment (Smyser, Page, and others, 2013). Six sites were used to 

assess experimental translocation or reintroduction of woodrats from other states. It was found that anthelmintic baiting 

facilitated woodrat reintroduction and recovery in treated sites 

as compared to untreated sites (Smyser, Johnson, and others, 

2013). Recently, an automated dispenser was developed for 

the delivery of anthelmintics or vaccines to raccoons in the 

wild (Smyser and others, 2015) and an anthelmintic fishmeal 

polymer bait was evaluated for the control of B. procyonis in 

wild raccoons (Smyser, Johnson, and others, 2015).

As promising as these results have been, because wild 

raccoons have high population densities in suburban areas and 

are likely to become reinfected after treatment, controlling 

Baylisascaris in raccoons in such areas by bait deworming 

may be challenging or ineffective, as well as expensive, if it 

is attempted on too large a scale. However, it may be advantageous to continually deworm a stable, localized resident 

population of raccoons on a regular basis as part of overall 

control efforts, combined with latrine cleanup and decontamination. It would also be very important to discourage people 

from intentionally feeding wild raccoons and to control other 

human-generated food sources (garbage, pet food) that serve 

to maintain large populations of raccoons (Roussere and 

others, 2003; Prange and others, 2003, 2004). This approach, 

combined with other population control methods (for example, 

contraception, neuter and release) could help stabilize and (or) 

limit local raccoon populations as well as reduce the establishment of new latrine sites. It could also reduce levels of 

raccoon fecal contamination and B. procyonis eggs in targeted 

areas, such as domestic or zoo environments, as well as habitats of threatened or endangered species, such as Allegheny 

woodrats.

Preventing Contact with Contaminated Areas 

or Articles

Captive raccoons, skunks, and other definitive hosts 

should be housed away from other species, in clean dedicated 

cages or enclosures that can be easily decontaminated. Regular 

feces removal, preferably daily but at least weekly, should be 

part of routine husbandry practices. These animals should not 

be housed in egg-contaminated enclosures or fed raw meat 

from wild animals (for example, rodents, rabbits, birds), as 

either could result in patent Baylisascaris infection and subsequent egg shedding (Kazacos and Boyce, 1989).

Once in the environment, Baylisascaris eggs can survive 

for years, so dealing with contaminated areas is problematic 

(Kazacos, 2001). Contaminated locations can be dealt with 

by using straightforward methods, but these must match the 

serious challenge put forth by the eggs. It is important to 
Disease Prevention and Control 77

prevent contact with known or suspected contaminated areas 

or articles until they can be properly assessed and effectively 

decontaminated, removed, and (or) destroyed (Kazacos, 

1983c, 2001). This would include cages or enclosures that 

had housed raccoons, skunks or other potential definitive 

hosts, raccoon latrine sites in and around the domestic or zoo 

environment, and other contaminated materials. It is important 

to prevent access by raccoons to buildings, animal facilities, 

and zoo exhibits and to prevent raccoon fecal contamination 

of hay, straw, and feed used for other animals. Fallen timber, 

large tree limbs, and rocks obtained from the wild and destined 

for use in animal enclosures or exhibits should first be carefully inspected, washed, and heat treated before use. It may be 

prudent to first strip the bark from dead trees that are collected 

for use in animal exhibits.

Education

Baylisascaris infection (NLM, OLM) can have devastating effects at many levels, affecting individuals, families, 

populations, facilities, and whole organizations. Baylisascaris

is not well known among the general public or the medical 

profession, but the situation has improved considerably in 

the last several decades. Despite this, a body of misinformation and misunderstanding exists concerning B. procyonis and 

its health effects and, justifiably so, the parasite continues to 

generate considerable fear in those who feel they may have 

been exposed or infected. Therefore, education of individuals 

and groups about the dangers associated with raccoons and 

Baylisascaris continues to be the most important aspect of 

prevention and control (Kazacos, 1991, 2000, 2001).

Educational programs about these parasites could be beneficial to a wide spectrum of people, including wildlife biologists, natural resources personnel, animal care directors and 

staff, wildlife rehabilitators, animal damage control officers, 

pet dealers and suppliers, public health personnel, veterinarians, physicians, and the general public. Questionnaire surveys 

in China (Xie and others, 2014; Y. Xie, written commun., 

2014) and the United States (Parkanzky, 2015) found a lack 

of awareness about B. procyonis, including its health effects, 

transmission, and relationship to animals, among people keeping raccoons and nonraccoon procyonids, including some zoo 

personnel. For parents and families, the main focus should 

be on infant children, because they are at the greatest risk of 

heavy infection and, therefore, demand close parental supervision to avoid becoming infected. Better availability of information about the real dangers posed to children by raccoons 

and Baylisascaris would be especially helpful for parents and 

pediatricians. It is critical that infectious disease specialists, 

neurologists (especially pediatric), and ophthalmologists are 

made aware of this common parasite and its potentially devastating consequences, and that this information is disseminated 

to the general medical community. Public awareness is also 

very important, so that parents can take appropriate precautions to protect their young children from infection. Parents 

who witness potential exposure of infant children at raccoon 

latrines must quickly seek correct information and prophylactic treatment from medical professionals, who should at least 

understand the seriousness of the situation or be able to obtain 

information quickly. Those seeking medical assistance because 

of neurologic disease and (or) problems with vision may need 

to communicate with their health professionals about this 

infection and its treatment, in addition to providing information about potential exposure to raccoons, skunks, or other 

animals and their feces.

Residential properties, including homes and yards, 

contiguous wooded areas, childcare facilities, farm buildings, 

etc., should be monitored for raccoon activity as well as latrine 

sites, and raccoons should be deterred from contaminating 

peridomestic areas and other facilities where children may 

have contact. Raccoon latrine sites should be identified and 

children and animals kept away from these areas until they can 

be decontaminated. Children should be taught at an early age 

not to play with animal feces, to identify and avoid raccoon 

latrines they may encounter in their environment, and to wash 

their hands after playing outside or with animals. Infection and 

clinical disease caused by Baylisascaris are both preventable 

through simple, straightforward means. These preventative 

measures may be most likely undertaken when people have 

greater awareness, understanding, and appreciation of the 

problem.

Treatment

Intestinal Worms 

Fecal samples from potential hosts kept in captivity, 

which could include raccoons, skunks, and even dogs, should 

be examined regularly by fecal flotation, and potentially 

infected animals should be strategically dewormed for Baylisascaris to prevent or decrease environmental contamination with eggs and possible transmission to humans and other 

animals (Kazacos and Boyce, 1989; Kazacos, 2001; Kazacos 

and others, 2013). Newly acquired raccoons and other hosts 

should be quarantined and dewormed immediately, with at 

least two additional anthelmintic treatments at 14-day intervals 

to ensure elimination of all developing worms (Kazacos and 

Boyce, 1989; Kazacos, 2001). Young raccoons, skunks and 

other definitive hosts, including dogs, pose a greater threat 

than adults, because they have a higher prevalence of infection and are often acquired during the prepatent period, when 

they will be harboring young developing worms but will be 

false negative for eggs (Kazacos, 1983b; Kazacos and Boyce, 

1989). These animals can be fecal negative for as long as 9–10 

weeks, then suddenly begin shedding large numbers of eggs 

when the worms mature, resulting in extensive contamination. Because the eggs are microscopic, shedding will not 

be noted by the owners, and when the eggs reach infectivity, the situation will become dangerous. Therefore, strategic 

deworming of young animals should be started early, at about 

5–6 weeks of age (and earlier for puppies, at 2 weeks of age, 
78 Baylisascaris Larva Migrans

due to transplacental transmission) and repeated regularly, 

for example, every 2 weeks for five or six treatments, then 

continued monthly (Kazacos and Boyce, 1989; Bauer and Gey, 

1995; Kazacos, 1991, 2001). The minimum known prepatent 

period of Baylisascaris in raccoons and skunks is about 1 

month postinfection (32 days for B. procyonis, via larvae), so 

once infections are eliminated, regular monthly deworming 

should prevent any future environmental contamination with 

eggs (Kazacos and Boyce, 1989; Bauer and Gey, 1995; Kazacos, 2001). Drugs effective against intestinal Baylisascaris 

include pyrantel pamoate, piperazine, fenbendazole, milbemycin oxime, moxidectin, and several others, all of which can be 

administered mixed in moist cat food, which is highly palatable to raccoons and skunks (Kazacos, 1986, 2001; Bauer and 

Gey, 1995; Bowman and others, 2005).

Migrating Larvae

As opposed to the treatment of intestinal worms, which is 

relatively easy, treatment of migrating Baylisascaris larvae is 

difficult and, with or without treatment, NLM due to Baylisascaris carries a guarded prognosis. The clinical efficacy of 

anthelmintic treatment of NLM depends on several factors, 

including drug pharmacokinetics, activity against larvae in 

the CNS, the level and duration of CNS infection, and the 

extent of CNS damage at the time of treatment (Kazacos, 

2001). Treatment of low-level or early CNS infection appears 

possible by using larvicidal drugs, such as albendazole, which 

effectively cross the blood-brain barrier and steroids to control 

inflammation (Murray and Kazacos, 2004; Gavin and others, 

2005; Kazacos and others, 2013). With early and aggressive 

therapy, cases of clinical NLM in humans and animals have 

been stabilized, and in some cases the patients have improved 

(Kazacos, 2001; Pai and others, 2007; Hajek and others, 2009; 

Jimenez Martinez and others, 2015; Langelier and others, written commun., 2015); however, in most reported cases treatment did not affect the progression of CNS disease.

It is very important that any treatments for Baylisascaris

NLM be started as early as possible, in an effort to kill larvae 

in the CNS and thus limit further damage and to kill larvae 

migrating in other tissues before they can reach the CNS. 

The anthelmintic of choice for treatment of Baylisascaris

NLM is albendazole, although diethylcarbamazine also had 

good efficacy. Ivermectin is not a good choice for clinical 

cases, because it does not readily cross the blood-brain barrier 

(Kazacos, 2001). Mice infected with fatal doses of B. procyonis eggs and treated daily with albendazole (25–50 mg/kg) for 

10 days starting 1–3 days postinfection were protected 95–100 

percent from developing CNS disease (Miyashita, 1993; Garrison, 1996). In cases of known or probable exposure, such as 

when a child is seen or suspected of ingesting material from a 

raccoon latrine, immediate treatment with albendazole daily 

for 10 days is recommended to prevent the development of 

NLM and CNS disease.

Preventive Treatment

In addition to treatment of early infection or clinical 

NLM, several anthelmintics show great promise as preventives 

for this infection in animals and possibly humans (Kazacos, 

2001). These include the pyrantel salts, pyrantel tartrate and 

pyrantel pamoate, which prevent initial infection in the intestine following egg ingestion, and thus subsequent larval migration and development of NLM. These drugs must be given 

daily or continuously to have a preventive effect against larvae 

hatching in the gut and are ineffective against larvae already 

in migration. Mice experimentally infected with B. procyonis

eggs and given pyrantel tartrate at 0.25 percent or 0.5 percent, 

or pyrantel pamoate at 0.2 percent concentration in their feed 

were fully protected against B. procyonis infection and NLM, 

which proved 100 percent fatal to untreated mice (Lindquist, 

1978). In cases of known or suspected exposure of animals to 

Baylisascaris eggs, pyrantel tartrate feed additive for swine 

(such as Banminth®48) or the pelleted formulation for horses 

(such as Strongid®C) can be mixed directly into the feed for 

large flightless birds, nonhuman primates, or other animals, 

or used as a top dressing on the animals’ usual food (Suedmeyer and others, 1996; Kazacos, 2001). Because of their high 

efficacy, acceptance, safety, and ease of use, these drugs are 

recommended for the prevention of Baylisascaris larval infection in mammals and birds (Kazacos, 2001). They are particularly useful on premises with ongoing problems, where the 

source of infection either has not been identified or effectively 

decontaminated. They would most likely be highly effective 

for preventing Baylisascaris infection in children, when given 

as a daily dose, but this has not been studied. Although pyrantel pamoate is a safe and approved human drug for general 

deworming, it is not labeled for prevention of Baylisascaris, 

even though it still could be used for this purpose.
Disease Prevention and Control 79

Decontamination Procedures for Baylisascaris Eggs

Baylisascaris eggs are exceptionally resistant to 

physical and chemical factors, but they can be effectively 

killed by using various forms of heat. In addition, they, 

along with raccoon feces, can be physically removed, 

making it possible to decontaminate contaminated areas 

and articles (Kazacos and Boyce, 1989; Kazacos, 2001). 

Guidelines for cleaning up raccoon latrines are on the Web 

sites of several governmental agencies. Baylisascaris eggs 

are resistant to all common disinfectants, including bleach, 

although certain solvent mixtures will kill them (Kazacos 

and Boyce, 1989; Kazacos, 2001). Treatment of an area 

with a solution of 20 percent bleach (1 percent sodium 

hypochlorite) removes the outer protein coat of the eggs, 

making them unable to adhere to surfaces and able to be 

washed away, but this will not kill them. B. procyonis eggs 

exposed to undiluted household bleach for 90 minutes 

were still viable (Shafir and others, 2011). Chemical treatments that will kill the eggs are generally not practical for 

use in the environment, and indeed may be illegal to use 

(Kazacos and Boyce, 1989; Kazacos, 1991, 2001).

Similar to other ascarid eggs, the thermal death point 

of B. procyonis eggs is about 144 degrees Fahrenheit (°F; 

62 degrees Celsius [°C]) (Shafir and others, 2007, 2011), 

making them susceptible to killing at temperatures lower 

than most people realize, and because heat comes in many 

forms, the eggs can be killed by various means, depending on the situation and location of contamination. Boiling 

or scalding water (150 °F to more than 160 °F), a steam 

cleaner, propane flame gun, autoclave, burning straw, or 

other means can be used to effectively decontaminate 

small or large areas of contaminated soil or concrete, metal 

cages, enclosures, holding pens, and contaminated tools 

and utensils (Kazacos and Boyce, 1989; Kazacos, 1991, 

2001). 

Government Web sites with guidelines for cleaning 

raccoon latrines.

Agency Website

Centers for Disease 

Control

http://www.cdc.gov/parasites/

baylisascaris/resources/

raccoonlatrines.pdf 

Santa Barbara County 

Animal Services

http://www.dshs.state.tx.us/idcu/

health/zoonosis/outdoor/

raccoonLatrine.pdf

Seattle and King County 

Public Health

http://www.kingcounty.gov/

healthservices/health/ehs/

RaccoonLatrine.aspx

A. Propane flame gun (VT 3–30 Red Dragon Vapor Torch) with 20-pound gas 

tank. (Photo by Kevin Kazacos and Sam Royer)

B. Use of propane flame gun to decontaminate a raccoon trap for 

B. procyonis eggs. (Photo by Sam Royer)

Direct flame from a propane gun is very effective for 

destroying eggs and is easily used to decontaminate live 

traps, cages, and enclosures that have held Baylisascarisinfected raccoons as well as shovels and other metal tools 

(A, B). This method is also used to decontaminate concretefloored animal rooms, kennel runs, and raccoon latrine sites 

in zoos and around homes (Pegg, 1977; Abdelrasoul and 

Fowler, 1979; Kazacos and Boyce, 1989; Kazacos, 1991, 

2001). Surface soil can be flamed, broken up, and turned 

over several times with a shovel or rake, and reflamed each 

time to ensure decontamination. Large tractor-mounted 

units are also available for use on soil areas. Obviously, 

appropriate precautions should be taken when using this 

method, particularly in or around buildings and other flammable materials, or another less dangerous heat method 

such as steam cleaning should be used instead. A wide 

variety of steam cleaners, both large and hand-held, are 

available and could be effectively used, depending on the 

situation.

Raccoons defecating on patios, porches, and decks 

can be dealt with by trapping and removing them and (or) 

regularly cleaning up their feces and using heat decontamination. Eggs in fresh feces are not yet dangerous and such 

material can be picked up weekly, bagged and disposed of in 

the trash, and the area treated with boiling or scalding water 

(for example, from a large pan or teapot) to kill any residual 

eggs that may be present (C). Heavily contaminated areas 

may also be dealt with by removing and discarding the top 

several inches of soil or substrate and replacing it; this may 

be combined with heat treatment of the area (D, E).
80 Baylisascaris Larva Migrans

Dried raccoon feces and other contaminated material 

(hay, straw, leaves) in attics, barn lofts, garages, etc., should 

be carefully removed, double-bagged, and disposed of in 

a landfill or destroyed by incineration. Lightly wetting the 

material first with water from a pump sprayer will allow 

removal without stirring up much dust. Residual material 

in buildings can then be removed by wiping the area with 

hot soapy water and (or) by using a powerful canister-type 

vacuum cleaner containing a fine (less than 15 micrometers) 

disposable filter bag. The surfaces can be further treated with 

steam or scalding water. Residential and commercial steam 

vacuums could also be used. Some commercial remediation 

companies have powerful truck-mounted vacuum systems that are very effective for removing and trapping fine 

particles including these eggs. Because Baylisascaris eggs 

die from desiccation in hot, dry attics and barn lofts, once 

raccoons have been prevented from regaining access to such 

areas, feces can be allowed to sit for an extended period, 

E. Daycare facility after remediation that included complete 

substrate removal. (Photo by William J. Murray)

C. Boiling water decontamination of a raccoon 

latrine area on a deck. (Photo by Kevin Kazacos)

D. Substrate removal during remediation of daycare facility 

in southern California following infection of children with 

B. procyonis from rooftop latrine. (Photo by William J. Murray)

Decontamination Procedures for Baylisascaris Eggs

especially during the hot summer, prior to cleanup. Recent 

studies found that it takes 7 months for B. procyonis eggs to 

die from desiccation (Shafir and others, 2011). The material 

can then be safely removed with few precautions because 

viable eggs will not be present. It is also theoretically possible to kill the eggs on surfaces using cryogenic freezing, 

similar to methods used for bed bugs and other pests, but 

this has not been examined.

Homeowners and professionals alike cleaning contaminated areas should wear personal protective equipment 

including disposable coveralls or old clothes, strong rubber 

gloves (for example, disposable nitrile), washable rubber 

boots, and a particulate face mask (for example, N–95) or 

better respiratory protection to prevent ingesting any eggs or 

inhaling fecal bacteria and fungi stirred up in dust (Kazacos 

and Boyce, 1989; Kazacos, 2001). When finished, disposable items should be bagged and discarded, incinerated, or 

otherwise properly disposed of.

The presence of eggs in soil or environmental debris, as 

well as the effectiveness of their destruction or removal, can 

be evaluated using routine centrifugal sedimentation-flotation methods (Kazacos, 1983c). It is very important that this 

be done by a trained parasitologist or microbiologist with 

the necessary experience and expertise for proper identification of Baylisascaris eggs. When present in environmental 

samples, the eggs must be separated from myriad pseudoparasites and other materials (pollen, other plant material, 

fungal spores, free-living invertebrates, other parasite eggs, 

etc.) that will also be recovered. For example, certain plant 

pollen bears a striking resemblance to Baylisascaris eggs 

and could easily be misidentified as such.
Coping with Baylisascaris Neural Larva Migrans—From 

Individual People, to Wildlife Populations, to Organizations

Disease Prevention and Control 81

The consequences of Baylisascaris infection must 

be dealt with at many levels, ranging from individual 

patients and their families to populations, facilities, and 

organizations.

Individuals and Families

For the individual patient, severe neural larva migrans 

(NLM) is life-changing as it can result in considerable 

brain damage from trauma and inflammation. Because 

there is a lag time in the development of clinical signs, by 

the time they are manifest and the diagnosis and treatment 

are considered, significant brain damage may already have 

occurred (Kazacos, 2000, 2001; Murray and Kazacos, 2004; 

Gavin and others, 2005; Kazacos and others, 2013; Singaravelu and others, 2016). This damage is often irreversible. The extent of the damage will vary depending on the 

infecting dose; however, there may be little hope for much 

improvement despite aggressive therapy. Thus, prevention 

of infection is vitally important with this parasite. When 

clinical signs indicate that a patient may have become 

infected, diagnosis and treatment must be initiated as early 

as possible. Once inflammation is brought under control and 

larvae are either killed or have become walled off, a patient 

may be stabilized albeit with varying neurologic deficits. In 

some cases of lower level infection in children, a patient’s 

condition may gradually improve as the brain continues 

to develop and neural pathways are developed and (or) 

“rewired” (Sanders, 2015). However, patients may continue 

to degenerate further as the brain undergoes postinflammatory atrophy and a significant loss of substance. Patients 

with severe NLM may end up in a vegetative state or with 

severe neurologic impairments: wheelchair-bound, incontinent, with loss of most normal functions, requiring nursing 

care for the rest of their lives. Because of severe neurologic 

disease, they may develop other disabilities, such as muscle 

atrophy in the extremities, severe tendon contractures that 

may require surgery, breathing difficulties and (or) aspiration pneumonia.

Families are changed forever as a once ebullient child 

undergoes a major change and downward spiral, being 

lost to the devastating effects of larva migrans encephalitis 

(Sanders, 2015). Parents and siblings must then adapt to 

an altered lifestyle that includes the demanding needs of 

the affected child (Huff and others, 1984; Park and others, 

2000; Rowley and others, 2000; Moertel and others, 2001; 

Reilly, 2008; Sanders, 2015). These challenges may be 

met with love and acceptance or may tear the family apart 

(Cheney, 2005; Reilly, 2008; Sanders, 2015). A parent may 

suffer from a level of guilt or blame over a lack of prior 

knowledge of the parasite or associated risks, as well as 

possible prevention of the infection in the affected loved 

one. The real tragedy is that baylisascariasis is a preventable 

condition based on education, greater awareness, and simple 

precautions (Kazacos, 2000; Murray and Kazacos, 2004; 

Gavin and others, 2005). 

Baylisascaris encephalitis has also occurred in numerous pets (Appendix 2). It is also important to note that the 

loss of pet animals, often considered valued members of the 

family, can have similarly devastating effects on the wellbeing of that family, and many pets have been lost to this 

disease. Owners are often guilt ridden, thinking they could 

have avoided the death of the pet somehow. In one particularly touching case, a single man in Indiana lost what was 

his entire family, namely ten parrots and conures, to severe 

neurologic disease in a short period of time. The birds 

had as many as 150 larvae in the brain alone. The owner 

was absolutely devastated by this and blamed himself for 

the loss of his birds, although it was purely an accident. 

Raccoons had accessed a seed mixture he had stored in his 

garage, and began defecating in the food container. Once 

the B. procyonis eggs became infective, he was essentially 

feeding highly contaminated feed to the birds, which all 

died in a short period of time (A.M. Lennox and others, 

unpub. data, 1996; Lennox and others, 2015).

Wildlife Populations

Early research by Tiner (1954) estimated that 

B. procyonis was responsible for 5 percent of natural rodent 

mortality in Illinois woodlots where raccoons were common. Even higher levels of infection and potential mortality 

were noted more recently in Indiana (Page, 1998; Page and 

others, 2001b,c). This is a significant population mortality 

level due to a single parasitic agent, and we now know that 

this very nonspecific parasite is affecting many more species than were ever considered earlier. Most cases of NLM 

in wildlife go undetected by humans due to their cryptic 

occurrence, isolated location, the small size of affected 

animals, and the fact that affected individuals are removed 

by predators and scavengers (Kazacos, 2001). For example, 

thousands of wild rodents and rabbits with neurologic 

disease are submitted to diagnostic laboratories or public 

health laboratories every year in the United States and 
Coping with Baylisascaris Neural Larva Migrans—From 

Individual People, to Wildlife Populations, to Organizations

82 Baylisascaris Larva Migrans

Canada with the primary concern being rabies; 99 percent 

of these animals are rabies negative (Fitzpatrick and others, 

2014) and, when examined, Baylisascaris encephalitis turns 

out to be a common cause of CNS disease in these and other 

animals (Richter and Kradel, 1964; Fleming and Caslick, 

1978; Kazacos, Appel, and Thacker, 1981; Roth and others, 

1982; Kazacos, 2001).

The real-time devastating effects of B. procyonis on 

particular animal populations are shown by the plight of the 

Allegheny woodrat, believed to have been extirpated from a 

large portion of its northeastern range by B. procyonis, and 

continuing to be adversely affected in other geographic areas 

(Kazacos, 2001; LoGiudice, 2003; Page and others, 2012) 

(see “The Plight of the Allegheny Woodrat”). In such cases, 

the interaction of raccoon populations with the local environment and other animal species becomes very important and 

is compounded by particular behaviors that place certain 

animal species at a greater risk of infection than others. Of 

particular importance is the potential effect of the parasite on 

threatened or endangered species that are already struggling 

due to human-generated effects on their environment or other 

factors. Even at low prevalence in raccoons and latrines, B. 

procyonis could still have important negative consequences, 

because a very small number of ingested eggs can result in 

fatal NLM in susceptible species (Page and others, 2012). 

One could easily extrapolate these findings (Tiner, 1954; 

Page and others, 2001b,c, 2012), combined with the widespread occurrence of NLM in mammals and birds, to indicate 

that B. procyonis is having a low-grade mortality effect on 

various animal populations within its range.

Facilities and Organizations

Baylisascaris NLM has had direct effects on agricultural 

operations, research animal facilities, zoos and wildlife parks, 

wildlife rehabilitation facilities, and human childcare facilities, based on significant outbreaks of neurologic disease in 

animals and humans. Numerous cases exist of either insidious or major devastating losses in animal operations (Appendix 3). In some cases, mortality was 100 percent or involved 

hundreds of animals.

Many zoos have ongoing problems with B. procyonis, because they are commonly overrun with raccoons at 

night, leading to widespread contamination of the facilities. 

Fatal or severe NLM has been seen in numerous zoo species. Serologic evidence of infection was found by ELISA 

in 76 of 259 (29 percent) nonhuman primates at 6 major 

zoos (Zimmerman and others, 2010), indicating just how 

commonly zoo animals are exposed to this parasite. In 

addition to raccoons, zoos may have problems with other 

nuisance wildlife species, including coyotes and skunks. An 

excellent example of this problem, which exists in many 

suburban zoos, occurred in the Los Angeles Zoo:

“The Los Angeles Zoo sits in the middle of Griffith 

Park, which is a large wild area. Previous nonmanagement 

of pests had allowed the zoo to become overrun with these 

animals, and problems had reached epidemic proportions 

in 1995. Coyotes living in the zoo were hunting gerenuk 

and flamingos, skunks were everywhere, and raccoons had 

free-roam of the zoo. In the past 3 years at the Los Angeles Zoo, we have seen numerous cases of central nervous 

system disease secondary to Baylisascaris” (Stringfield and 

Sedgwick, 1997).

These problems were met by an immediate and 

aggressive, multifaceted response following a change in 

zoo management. Response measures entailed trapping and 

removing the resident raccoon, skunk, and coyote populations from zoo property, repairing gaps in the perimeter fencing to prevent future influx, trimming trees and 

overhanging foliage, installing wire mesh barriers along 

the bottoms of exhibits to prevent animal access, repairing garbage bins, cleaning and rehabilitating contaminated 

exhibits, and instituting ongoing surveillance and control 

measures, coupled with educating staff. The scope of the 

task was daunting; however, with thoughtful planning and 

implementation the program was successful, and nuisance 

wildlife and Baylisascaris transmission to zoo animals 

were both brought under control.

Although most human cases have involved individual 

children, several have involved facilities where additional 

children or adults were exposed to infection. In one case 

in southern California, an 11-month-old boy at a well-run 

university childcare facility developed marked eosinophilic 

meningoencephalitis with blindness due to B. procyonis

(Schultz, 2002; Murray and Kazacos, 2004; K.R. Kazacos, 

unpub. data, 2002–4; Sanders, 2015). He was in a facility 

where over 80 other children in his class group, as well as 

others, were potentially exposed. Several other children 

also were seropositive, indicating infection, but without 

obvious clinical signs. Infection was linked to the presence 

of raccoon latrines on the sloped metal roof of the building, with feces and eggs washing down into the infant and 

toddler play area below (A–C). Following this outbreak, 

the facility was closed under public health quarantine and 
Coping with Baylisascaris Neural Larva Migrans—From 

Individual People, to Wildlife Populations, to Organizations

C. Infant and toddler play area in contaminated childcare 

facility in California. (Photo by William J. Murray)

Disease Prevention and Control 83

extensively remediated before being allowed to reopen 

and accept children again. The victim’s family moved to 

the northeastern United States, was able to secure a major 

monetary settlement for his care, and the child continues to 

improve following aggressive therapy at the time of infection and subsequent major physical and learning rehabilitation (K.R. Kazacos, unpub. data, 2004–15; Sanders, 2015) 

(D).

Cases also have involved adults in group homes or state 

facilities for developmentally disabled individuals. One case 

in southern California involved a 17-year-old boy admitted to a hospital emergency room in a coma, suffering from 

severe Baylisascaris encephalitis from which he later died 

(Kazacos, Gavin, and others, 2002). This patient lived in a 

well-run group home for developmentally disabled adolescents, where he displayed abnormal behavior including 

eating sand from the sandbox play area. This sandbox was 

the site of a raccoon latrine and had extensive contamination 

with B. procyonis eggs, and the property had to be decontaminated and remediated to prevent other cases. A similar 

case occurred in Oregon, where a 21-year-old man in a state 

facility became infected due to geophagia (Cunningham and 

others, 1994) and subsequently suffered a long course of 

worsening central nervous system problems and therapies, 

including partial cranial lobotomy. These cases indicate that 

individuals in group facilities may be at a particular risk of 

infection due to their developmentally delayed status and 

altered behavior, their young age, or both.

Besides the devastating health effects on individuals, 

these incidents raise important questions concerning previously unconsidered legal liability issues involving 

B. procyonis infection in such facilities and, as previously 

mentioned, at least one case resulted in a major monetary 

settlement following legal redress. It is definitely in their 

own best interest, as well as that of their clients and endusers, that such organizations address potential issues with 

this parasite at their facilities, through education, heightened awareness and preventive measures, increased animal 

control and exclusion, fecal cleanup, sanitation, surveillance, 

and other routine precautions. 

A. (left) Raccoon latrine on the rooftop of 

a childcare facility in southern California 

associated with B. procyonis infection in 

children. (Photo by William J. Murray)

B. (above) Closeup of raccoon latrine on 

rooftop of childcare facility in California. 

(Photo by William J. Murray)

D. Child infected with B. procyonis in childcare facility in California, 

several years later following treatment and extensive physical and 

learning rehabilitation. (Photos by Anthony R. and Kevin Kazacos)
84 Baylisascaris Larva Migrans

References Cited

Abdelrasoul, K., and Fowler, M., 1979, Epidemiology of 

ascarid infection in captive carnivores: Proceedings of the 

American Association of Zoo Veterinarians, p. 105–106a.

Agnew, D.W., Kazacos, K.R., Watson, G.L., Yamini, B., 

Barbiers, R., and Garrison, R.D., 1994, Neural larva 

migrans due to Baylisascaris procyonis in red kangaroos: 

Proceedings of the American Association of Veterinary 

Parasitologists, v. 39, p. 68.

Akao, N., Hayashi, E., Sato, H., Fujita, K., and Furuoka, H., 

2003, Diffuse retinochoroiditis due to Baylisascaris 

procyonis in Mongolian gerbils: Journal of Parasitology, 

v. 89, p. 174–175.

Akao, N., and Ohta, N., 2007, Toxocariasis in Japan: 

Parasitology International, v. 56, p. 87–93.

Aliev, F.F., and Sanderson, G.C., 1966, Distribution and status 

of the raccoon in the Soviet Union: Journal of Wildlife 

Management, v. 30, p. 497–502.

Al-Lebban, H.M., and Lindquist, W.D., 1979, Comparison 

between migratory behavior of Toxocara canis and 

Baylisascaris procyonis larvae in orally and parenterally 

infected mice: Proceedings of the Conference of Research 

Workers in Animal Disease, Abstract 130.

Amaro, M.H., 2000, Reports of diffuse unilateral subacute 

neuroretinitis: Archives of Ophthalmology, v. 118, p. 1593.

American Ornithologists’ Union, 2012, Check-list of North 

American birds: http://checklist.aou.org/taxa/.

Amundson, T., and Marquenski, S., 1986, Baylisascaris procyonis in Wisconsin raccoons: Madison, Wisconsin 

Department of Natural Resources, p. 1–7.

Anderson, B.C., 1999, Congenital Baylisascaris sp. larval 

migrans in a newborn lamb: Journal of Parasitology, v. 85, 

p. 128–129.

Anderson, D.C., Greenwood, R., Fishman, M., and Kagan, 

I.G., 1975, Acute infantile hemiplegia with cerebrospinal 

fluid eosinophilic pleocytosis: An unusual case of visceral 

larva migrans: Journal of Pediatrics, v. 86, p. 247–249.

Anderson, S.B., and Mills, J., 1991, Occurrence of the raccoon 

roundworm (Baylisascaris procyonis) in raccoons in 

Nova Scotia: Kentville, Nova Scotia Department of 

Lands and Forests, Technical Note No. 67, 2 p.

Armstrong, D.L., Montali, R.J., Kazacos, K.R., and Doster, 

A.R., 1987, Cerebrospinal nematodiasis in blue and gold 

macaws and scarlet macaws associated with Baylisascaris 

procyonis: Proceedings of the First International Conference 

on Zoological and Avian Medicine, p. 489–490.

Armstrong, D.L., Montali, R.J., Doster, A.R., and Kazacos, 

K.R., 1989, Cerebrospinal nematodiasis in macaws due to 

Baylisascaris procyonis: Journal of Zoo and Wildlife 

Medicine, v. 20, p. 354–359.

Averbeck, G.A., Vanek, J.A. Stromberg, B.E. and Laursen, 

J.R., 1995, Differentiation of Baylisascaris species, Toxocara canis, and Toxascaris leonina infections in dogs: 

Compendium on Continuing Education for the Practicing 

Veterinarian, v. 17, p. 475–478, 511.

Babero, B.B., and Shepperson, J.R. 1958, Some helminths of 

raccoons in Georgia: Journal of Parasitology, v. 44, p. 519.

Bafundo, K.W., Wilhelm, W.E., and Kennedy, M.L., 1980, 

Geographic variation in helminth parasites from the digestive tract of Tennessee raccoons, Procyon lotor: Journal of 

Parasitology, v. 66, p. 134–139.

Balcom, B.J., and Yahner, R.H., 1996, Microhabitat and landscape characteristics associated with the threatened Allegheny woodrat: Conservation Biology, v. 10, p. 515–525.

Ball, R.L., Dryden, M., Wilson, S., and Veatch, J., 1998, 

Cerebrospinal nematodiasis in a white-handed gibbon 

(Hylobates lar) due to Baylisascaris sp: Journal of Zoo and 

Wildlife Medicine, v. 29, p. 221–224.

Barnstable, R.W., and Dyer, W.G., 1974, Gastrointestinal helminths of the raccoon, Procyon lotor, in southern Illinois: 

Transactions of the Illinois State Academy of Science, v. 67, 

p. 451–460.

Bartoszewicz, M., 2011, NOBANIS—Invasive Alien Species 

Fact Sheet—Procyon lotor: Online database of the European Network on Invasive Alien Species— NOBANIS, S 

accessed May 25, 2104, at http://www.nobanis.org/files/

factsheets/Procyon_lotor.pdf.

Bartoszewicz, M., Okarma, H., Zalewski, A., and Szczęsna, J., 

2008, Ecology of the raccoon (Procyon lotor) from western 

Poland: Annales Zoologici Fennici, v. 45, p. 291–298.

Bauer, C., and Gey, A., 1995, Efficacy of six anthelmintics 

against luminal stages of Baylisascaris procyonis in naturally infected raccoons (Procyon lotor): Veterinary Parasitology, v. 60, p. 155–159.

Beasley, J.C., Eagan, T.S., II, Page, L.K., Hennessy, C.A., and 

Rhodes, O.E., Jr., 2013, Baylisascaris procyonis infection 

in white-footed mice: predicting patterns of infection from 

landscape habitat attributes: Journal of Parasitology, v. 99, 

p. 743–747.

Beaver, P.C., 1969, The nature of visceral larva migrans: Journal of Parasitology, v. 55, p. 3–12.
References Cited 85

Beck, S.E., Kelly, K., Mankowski, J., Nolte, J., and Graham, 

D., 2010, Case 8: 1 year, 4 month-old female titi monkey 

(62078): Proceedings of the 2010 Primate Pathology Workshop, ACVP/ASVCP Pre-Meeting Workshop of American 

College of Veterinary Pathologists, Baltimore, Maryland, 

October 30, 2010, p. 26–29.

Beltrán-Beck, B., Garcia, F.J., and Gortazar, C., 2012, Raccoons in Europe: disease hazards due to the establishment of an invasive species: European Journal of Wildlife 

Research, v. 58, no. 1, p. 5–15.

Berry, J.F., 1985, Phylogenetic relationship between Baylisascaris spp. Sprent, 1968 (Nematoda: Ascarididae) from 

skunks, raccoons and groundhogs in southern Ontario: 

Guelph, Ontario, University of Guelph, M.S. Thesis, 99 p.

Birch, G.L., Feldhamer, G.A., and Dyer, W.G., 1994. Helminths of the gastrointestinal tract of raccoons in southern 

Illinois with management implications of Baylisascaris 

procyonis occurrence: Transactions of the Illinois State 

Academy of Science, v. 87, p. 165–170.

Blizzard, E.L., Davis, C.D., Henke, S., Long, D.B., Hall, 

C.A., and Yabsley, M.J., 2010, Distribution, prevalence, 

and genetic characterization of Baylisascaris procyonis in 

selected areas of Georgia: Journal of Parasitology, v. 96, 

p. 1128–1133.

Blizzard, E.L., Yabsley, M.J., Beck, M.F., and Harsch, S., 

2010, Geographic expansion of Baylisascaris procyonis 

roundworms, Florida, USA: Emerging Infectious Diseases, 

v. 16, p. 1803–1804.

Bluett, R.D., Hubert, G.F., Jr., and Miller, C.A., 2003, Regulatory oversight and activities of wildlife control operators in 

Illinois: Wildlife Society Bulletin, v. 31, p. 104–116.

Boddicker, M.L., and Progulske, D.R., 1968, Helminth 

parasites of raccoon in South Dakota: Proceedings of the 

South Dakota Academy of Science, v. 47, p. 161–166.

Boggess, E.K., 1994, Raccoons, in Hygnstrom, S.E., Timm, 

R.M., and Larson, G.E., Prevention and control of wildlife 

damage: University of Nebraska-Lincoln, p. C–101–C–107.

Boschetti, A., and Kasznica, J., 1995, Visceral larva migrans 

induced eosinophilic cardiac pseudotumor: A cause of sudden death in a child: Journal of Forensic Sciences, v. 40, 

p. 1097–1099.

Bowman, D.D., 1987, Diagnostic morphology of four larval 

ascaridoid nematodes that may cause visceral larva migrans: 

Toxascaris leonina, Baylisascaris procyonis, Lagochilascaris sprenti, and Hexametra leidyi: Journal of Parasitology, 

v. 73, p. 1198–1215.

Bowman, D.D., Ulrich, M.A., Gregory, D.E., Neumann, N.R., 

Legg, W., and Stansfield, D., 2005, Treatment of Baylisascaris procyonis infections in dogs with milbemycin oxime: 

Veterinary Parasitology, v. 129, p. 285–290.

Boyce, W.M., Branstetter, B.A., and Kazacos, K.R., 1988a, In 

vitro culture of Baylisascaris procyonis and initial analysis 

of larval excretory-secretory antigens: Proceedings of the 

Helminthological Society of Washington, v. 55, p. 15–18.

Boyce, W.M., Branstetter, B.A., and Kazacos, K.R., 1988b, 

Comparative analysis of larval excretory-secretory antigens 

of Baylisascaris procyonis, Toxocara canis and Ascaris 

suum by western blotting and enzyme immunoassay: International Journal for Parasitology, v. 18, p. 109–113.

Boyce, W.M., Asai, D.J., Wilder, J.K., and Kazacos, K.R., 

1989, Physicochemical characterization and monoclonal and polyclonal antibody recognition of Baylisascaris 

procyonis larval excretory-secretory antigens: Journal of 

Parasitology 75, p. 540–548.

Brasil, O.F.M., Lewis, H., and Lowder, C.Y., 2006, Migration 

of Baylisascaris procyonis into the vitreous: British Journal 

of Ophthalmology, v. 90, p. 1203–1204.

Brinch, K.S., 2006, Baylisascaris procyonis (vaskebjornens 

spolorm) hos danske vaskebjorne: Dansk Veterinaertidsskrift, v. 89, p. 28–35.

Brinkman, W.B., Kazacos, K.R., Gavin, P.J., Binns, H.J., 

Robichaud, J.D., O’Gorman, M., and Shulman, S.T., 2003, 

Seroprevalence of Baylisascaris procyonis (raccoon roundworm) in Chicago area children: Proceedings of the Annual 

Meeting of the Pediatric Academic Societies, Abstract 1872.

Brown, L.W., and Huff, D., 1983, Human disease due to the 

raccoon ascarid Baylisascaris procyonis: a new cause of 

eosinophilic meningitis: Proceedings of the Child Neurology Society, Abstract 110.

Campbell, G.A., Hoover, J.P., Russell, W.C., and Breazile, 

J.E., 1997, Naturally occurring cerebral nematodiasis due 

to Baylisascaris larval migration in two black-and-white 

ruffed lemurs (Varecia variegata variegata) and suspected 

cases in three emus (Dromaius novaehollandiae): Journal 

of Zoo and Wildlife Medicine, v. 28, p. 204–207.

Carlson, M.S., and Nielsen, S.W., 1984, Jejunal obstruction 

due to Baylisascaris procyonis in a raccoon: Journal of 

the American Veterinary Medical Association, v. 185, 

p. 1396–1397.

Casella, A.M.B., Farah, M.E., and Belfort, R., Jr., 1998, 

Anthelmintic drugs in diffuse unilateral subacute 

neuroretinitis: American Journal of Ophthalmology, 

v. 125, p. 109–111.
86 Baylisascaris Larva Migrans

Centers for Disease Control and Prevention, 2015, Bringing an 

animal into the United States: accessed September 10, 2015, 

at http://www.cdc.gov/importation/bringing-an-animal-intothe-United-States/index.html.

Chandler, A.C., 1942, The helminths of raccoons in east 

Texas: Journal of Parasitology, v. 28, p. 255–268.

Chavez, D.J., LeVan, I.K., Miller, M.W., and Ballweber, L.R., 

2012, Baylisascaris procyonis in raccoons (Procyon lotor) 

from eastern Colorado, an area of undefined prevalence: 

Veterinary Parasitology, v. 185, p. 330–334.

Cheney, P., 2005, The danger that lurks in the backyard dirt: 

The Globe and Mail, Globe Toronto, Saturday, 

October 8, 2005, M1.

Ching, H.L., Leighton, B.J., and Stephen, C., 2000, Intestinal 

parasites of raccoons (Procyon lotor) from southwest British Columbia: Canadian Journal of Veterinary Research, 

v. 64, p. 107–111.

Chris, A., 2005, Baylisascaris–Canada (Ontario): ProMEDmail 2005; 6 September: 20050907.2660 (http://www.

promedmail.org).

Chun, C.S., Kazacos, K.R., Glaser, C., Bardo, D., Dangoudoubiyam, S., and Nash, R., 2009, Global neurological deficits 

with Baylisascaris encephalitis in a previously healthy 

teenager: Pediatric Infectious Disease Journal, v. 28, 

p. 925–927.

Church, E.M., Wyand, D.S., and Lein, D.H., 1975, Experimentally induced cerebrospinal nematodiasis in rabbits 

(Oryctolagus cuniculus): American Journal of Veterinary 

Research, v. 36, p. 331–335.

Cialdini, A.P., de Souza, E.C., and Avila, M.P., 1999, The first 

South American case of diffuse unilateral subacute neuroretinitis caused by a large nematode: Archives of Ophthalmology, v. 117, p. 1431–1432.

Ciarlini, P., Gutierrez, Y., Gambetti, P., and Cohen, M., 

2011, Necrotizing eosinophilic meningoencephalitis: Case 

2011–06, 52nd Annual Diagnostic Slide Session, 87th Annual 

Meeting of the American Association of Neuropathologists, 

Seattle, Washington, June 23–26, 2011, accessed 

July 30, 2014, at http://neuro.pathology.pitt.edu/DSSFiles/

DSSCases2011.htm.

Clark, G.M., Philip, C.B., and Fadness, L., 1969, Observations on the effect in mice of concurrent infections with 

migratory Ascaris columnaris larvae and certain neurotropic 

arboviruses: Folia Parasitologica, v. 16, p. 67–73.

Coates, J.W., Siegert, J., Bowes, V.A., and Steer, D.G., 1995, 

Encephalitic nematodiasis in a Douglas squirrel and a rock 

dove ascribed to Baylisascaris procyonis: Canadian Veterinary Journal, v. 36, p. 566–569.

Cole, R.A., and Shoop, W.L., 1987, Helminths of the raccoon 

(Procyon lotor) in western Kentucky: Journal of Parasitology, v. 73, p. 762–768.

Conboy, G., 1996, Diagnostic parasitology: Canadian Veterinary Journal, v. 37, p. 181–182.

Conboy, G., Stewart, T.A., and Taylor, A., 2010, Baylisascaris 

procyonis infection in raccoons and dogs on Prince Edward 

Island, Canada: Proceedings of the American Association of 

Veterinary Parasitologists, v. 55, p. 56.

Conraths, F.J., Bauer, C., Cseke, J., and Laube, H., 1996, 

Arbeitsplatzbedingte Infektionen des Menschen mit dem 

Waschbärspulwurm Baylisascaris procyonis: Arbeitsmedizin Sozialmedizin Umweltmedizin, v. 31, p. 13–17. [In 

German with English summary.]

Cooney, T.A., 1989, Environmental contamination with 

Baylisascaris procyonis in an urban park: West Lafayette, 

Indiana, Purdue University, M.S. Thesis, 125 p.

Cortez, R., Denny, J.P., Muci-Mendoza, R., Ramirez, G., 

Fuenmayor, D., and Jaffe, G.J., 2005, Diffuse unilateral 

subacute neuroretinitis in Venezuela: Ophthalmology, 

v. 112, p. 2110–2114.

Cottrell, W.O., Heagy, R.L., Johnson, J.B., Marcantuno, R., 

and Nolan, T.J., 2014, Geographic and temporal prevalence 

of Baylisascaris procyonis in raccoons (Procyon lotor) in 

Pennsylvania, USA: Journal of Wildlife Diseases, v. 50, 

p. 923-927.

Craig, S.J., Conboy, G.A., and Hanna, P.E., 1995, Baylisascaris sp. infection in a guinea pig: Laboratory Animal Science, 

v. 45, p. 312–314.

Cranfield, M.R., Barker, I.K., Mehren, K.G., and Rapley, 

W.A., 1984, Canine distemper in wild raccoons (Procyon 

lotor) at the metropolitan Toronto zoo: Canadian Veterinary 

Journal, v. 25, p. 63–66.

Crosley, C.J., and Kazacos, K.R., 2005, Baylisascaris

meningo-encephalitis in three infants: Abstract P76, Proceedings of the 34th Annual Meeting of the Child Neurology Society, Los Angeles, California, Sept 28–Oct 1, 2005, 

Annals of Neurology, v. 58 (suppl. 9), p. S114.

Cunningham, C.K., Kazacos, K.R., McMillan, J.A., Lucas, 

J.A., McAuley, J.B., Wozniak, E.J., and Weiner, L.B., 1994, 

Diagnosis and management of Baylisascaris procyonis

infection in an infant with nonfatal meningoencephalitis: 

Clinical Infectious Diseases, v. 18, p. 868–872.

Dade, A.W., Williams, J.F., Whitenack, D.L., and Williams, 

C.S.F., 1975, An epizootic of cerebral nematodiasis in rabbits due to Ascaris columnaris: Laboratory Animal Science, 

v. 25, p. 65–69.
References Cited 87

Dade, A.W., Williams, J.F., Trapp, A.L., and Ball, W.H., Jr., 

1977, Cerebral nematodiasis in captive nutria: Journal of 

the American Veterinary Medical Association, v. 171, 

p. 885–886.

Dangoudoubiyam, S., and Kazacos, K.R., 2009, Differentiation of larva migrans caused by Baylisascaris procyonis and 

Toxocara species by Western blotting: Clinical and Vaccine 

Immunology, v. 16, p. 1563–1568.

Dangoudoubiyam, S., Vemulapalli, R., and Kazacos, K.R., 

2009, PCR assays for detection of Baylisascaris procyonis

eggs and larvae: Journal of Parasitology, v. 95, p. 571–577.

Dangoudoubiyam, S., Vemulapalli, R., Hancock, K., and 

Kazacos, K.R., 2010, Molecular cloning of an immunogenic 

protein of Baylisascaris procyonis and expression in 

Escherichia coli for use in developing improved serodiagnostic assays: Clinical and Vaccine Immunology, v. 17, 

p. 1933–1939.

Dangoudoubiyam, S., Vemulapalli, R., Ndao, M., and Kazacos, K.R., 2011, Recombinant antigen-based enzyme-linked 

immunosorbent assay for diagnosis of Baylisascaris procyonis larva migrans: Clinical and Vaccine Immunology, v. 18, 

p. 1650–1655.

Davidson, R.K., Øines, Ø., Hamnes, I.S., and Schulze, J.E., 

2013, Illegal wildlife imports more than just animals—Baylisascaris procyonis in raccoons (Procyon lotor) in Norway: 

Journal of Wildlife Diseases, v. 49, p. 986–990.

de Almeida, M.H., 1987, Nuisance furbearer damage control 

in urban and suburban areas, in Novak, M., Baker, J.A., 

Obbard, M.E., and Malloch, B., eds., Wild furbearer management and conservation in North America: North Bay, 

Canada, Ontario Trappers Association, p. 996–1006.

Deeb, B.J., and DiGiacomo, R.F., 1994, Cerebral larva 

migrans caused by Baylisascaris sp in pet rabbits: Journal 

of the American Veterinary Medical Association, v. 205, 

p. 1744–1747.

Diab, S.S., Uzal, F.A., Giannitti, F., and Shivaprasad, H.L., 

2012, Cerebrospinal nematodiasis outbreak in an urban 

outdoor aviary of cockatiels (Nymphicus hollandicus) in 

southern California: Journal of Veterinary Diagnostic 

Investigation, v. 24, p. 994–999.

Dixon, D., Reinhard, G.R., Kazacos, K.R., and Arriaga, C., 

1988, Cerebrospinal nematodiasis in prairie dogs from a 

research facility: Journal of the American Veterinary 

Medical Association, v. 193, p. 251–253.

Done, L.B., and Tamura, Y., 2014, A presumptive case of 

Baylisascaris procyonis in a feral green-cheeked Amazon 

parrot (Amazona viridigenalis): Journal of Zoo and Wildlife 

Medicine, v. 45, p. 157–160.

Donnelly, J.J., Sakla, A.A., Khatami, M., and Rockey, J.H., 

1989, Baylisascaris procyonis (Stefanski and Zarnowski, 

1951) Ascarididae: Nematoda. II. Third stage larvae, 

morphogenesis and migratory behaviour: Assiut Veterinary 

Medical Journal, v. 21, p. 77–85.

Dubey, J.P., 1982, Baylisascaris procyonis and eimerian infections in raccoons: Journal of the American Veterinary Medical Association, v. 181, p. 1292–1294.

Dubey, J.P., Hamir, A.N., Hanlon, C.A., and Rupprecht, C.E., 

1992, Prevalence of Toxoplasma gondii infection in raccoons: Journal of the American Veterinary Medical Association, v. 200, p. 534–536.

Eberhard, M.L., Nace, E.K., Won, K.Y., Punkosdy, G.A., 

Bishop, H.S., and Johnston, S.P., 2003, Baylisascaris procyonis in the metropolitan Atlanta area: Emerging Infectious 

Diseases, v. 9, p. 1636–1637.

Ermer, E.M., and Fodge, J.A., 1986, Occurrence of the raccoon roundworm in raccoons in western New York: New 

York Fish and Game Journal, v. 33, p. 58–61.

Evans, R.H., 2001, Baylisascaris procyonis (Nematoda: 

Ascaridae) in raccoons (Procyon lotor) in Orange County, 

California: Vector Borne Zoonotic Diseases, v. 1, 

p. 239–242.

Evans, R.H., 2002, Baylisascaris procyonis (Nematoda: 

Ascarididae) larva migrans in free-ranging wildlife in 

Orange County, California: Journal of Parasitology, v. 88, 

p. 299–301.

Evans, R.H., and Tangredi, B., 1985, Cerebrospinal nematodiasis in free-ranging birds: Journal of the American Veterinary Medical Association, v. 187, p. 1213–1214.

Fan, C.K., Holland, C.V., Loxton, K., and Barghouth, U., 

2015, Cerebral toxocariasis: Silent progression to neurodegenerative disorders?: Clinical Microbiology Reviews, v. 

28, p. 663–686. 

Feigley, H.P., 1992, The ecology of the raccoon in suburban 

Long Island, N.Y. and its relation to soil contamination with 

Baylisascaris procyonis ova: Syracuse, New York, State 

University of New York College of Environmental Science 

and Forestry, Ph.D. Thesis, 139 p.

Ferris, D.H., Lord, R.D., Jr., Beamer, P.D., and Fritz, T.E., 

1960, A new disease in Illinois cottontails: Journal of 

Wildlife Management, v. 24, p. 179–184.

Fitzgerald, S.D., White, M.R., and Kazacos, K.R., 1991, 

Encephalitis in two porcupines due to Baylisascaris larval 

migration: Journal of Veterinary Diagnostic Investigation, 

v. 3, p. 359–362.
88 Baylisascaris Larva Migrans

Fitzpatrick, J.L., Dyer, J.L., Blanton, J.D., Kuzmin, I.V., and 

Rupprecht, C.E., 2014, Rabies in rodents and lagomorphs 

in the United States, 1995–2010: Journal of the American 

Veterinary Medical Association, v. 245, p. 333–337.

Fleming, W.J., and Caslick, J.W., 1978, Rabies and cerebrospinal nematodiasis in woodchucks (Marmota monax) from 

New York: Cornell Veterinarian, v. 68, p. 391–395.

Fleming, W.J., Georgi, J.R., and Caslick, J.W., 1979, Parasites 

of the woodchuck (Marmota monax) in central New York: 

Proceedings of the Helminthological Society of Washington, v. 46, p. 115–127.

Forrester, D.J., 1992, Parasites and diseases of wild mammals 

in Florida: Gainesville, University Press of Florida, 480 p.

Fox, A.S., Kazacos, K.R., Gould, N.S., Heydemann, P.T., 

Thomas, C., and Boyer, K.M., 1985, Fatal eosinophilic 

meningoencephalitis and visceral larva migrans caused by 

the raccoon ascarid Baylisascaris procyonis: New England 

Journal of Medicine, v. 312, p. 1619–1623.

Franssen, F., Xie, K., Sprong, H., and van der Giessen, J., 

2013, Molecular analysis of Baylisascaris columnaris

revealed mitochondrial and nuclear polymorphisms: 

Parasites and Vectors, v. 6, p. 124–133.

Fritz, T.E., Smith, D.E., and Flynn, R.J., 1968, A central nervous system disorder in ground squirrels (Citellus tridecemlineatus) associated with visceral larva migrans: Journal of 

the American Veterinary Medical Association, v. 153, 

p. 841–844.

Furuoka, H., Sato, H., Kubo, M., Owaki, S., Kobayashi, Y., 

Matsui, T., and Kamiya, H., 2003, Neuropathological observation of rabbits (Oryctolagus cuniculus) affected with 

raccoon roundworm (Baylisascaris procyonis) larva 

migrans in Japan: Journal of Veterinary Medical Science, 

v. 65, p. 695–699.

García, J.T., García, F.J., Alda, F., González, J.L., Aramburu, 

M.J., Cortés, Y., Prieto, B., Pliego, B., Pérez, M., Herrera, 

J., and García-Román, L., 2012, Recent invasion and status 

of the raccoon (Procyon lotor) in Spain: Biological Invasions, v. 14, p. 1305–1310. 

Garlick, D.S., Marcus, L.C., Pokras, M., and Schelling, S.H., 

1996, Baylisascaris larva migrans in a spider monkey 

(Ateles sp.): Journal of Medical Primatology, v. 25, 

p. 133–136.

Garrison, R.D., 1996, Evaluation of anthelmintic and corticosteroid treatment in protecting mice (Mus musculus) from 

neural larva migrans due to Baylisascaris procyonis: West 

Lafayette, Indiana, Purdue University, M.S. Thesis, 102 p.

Garrison, R.D., and Kazacos, K.R., 1994, Effects of freezing on the development and infectivity of Baylisascaris 

procyonis eggs: Proceedings of the American Association of 

Veterinary Parasitologists, v. 39, p. 68.

Gass, J.D.M., 1997, Inflammatory diseases of the retina and 

choroid, in Stereoscopic atlas of macular diseases: diagnosis and treatment, 4th ed.: St. Louis, Missouri, Mosby-Year 

Book Inc., p. 622–628.

Gass, J.D.M., and Braunstein, R.A., 1983, Further observations concerning the diffuse unilateral subacute neuroretinitis syndrome: Archives of Ophthalmology, v. 101, 

p. 1689–1697.

Gass, J.D.M., Gilbert, W.R., Jr., Guerry, R.K., and Scelfo, R., 

1978, Diffuse unilateral subacute neuroretinitis: Ophthalmology, v. 85, p. 521–545.

Gass, J.D.M., Callanan, D.G., and Bowman, C.B., 1992, 

Oral therapy in diffuse unilateral subacute neuroretinitis: 

Archives of Ophthalmology, v. 110, p. 675–680.

Gatcombe, R.R., Narayanan, J., Dangoudoubiyam, S., Kazacos, K.R., and Hill, V., 2010, Evaluation of a molecular 

beacon real-time PCR assay for detection of Baylisascaris 

procyonis in different soil types and water samples: 

Parasitology Research, v. 106, p. 499–504.

Gavin, P.J., Kazacos, K.R., and Shulman, S.T., 2005, Baylisascariasis: Clinical Microbiology Reviews, v. 18, p. 703–718.

Gavin, P.J., Kazacos, K.R., Tan, T.Q., Brinkman, W.B., Byrd, 

S.E., Davis, A.T., Mets, M.B., and Shulman, S.T., 2002, 

Neural larva migrans caused by the raccoon roundworm 

Baylisascaris procyonis: Pediatric Infectious Disease Journal, v. 21, p. 971–975.

Gehrt, S.D., 2003, Raccoons (Procyon lotor) and allies, in

Feldhamer, G.A., Thompson, B.C., and Chapman, J.A., 

eds., Wild mammals of North America: Biology, management, and conservation, 2nd ed: Baltimore, Maryland, Johns 

Hopkins University Press, p. 611–634.

Geptner, V.G., and Naumov, N.P., 1967, Mammals of USSR, 

Vol. 2: Moscow, Visaja skola. [In Russian.]

Gey, A.B., 1998, Synopsis der Parasitenfauna des Waschbären 

(Procyon lotor) unter Berücksichtigung von Befunden aus 

Hessen: Giesen, Germany, Justus Liebig Universität, 

Dissertation, Fachbereich Veterinärmedizin, 203 p. 

[In German.]

Goldberg, M.A., Kazacos, K.R., Boyce, W.M., Ai, E., and 

Katz, B., 1993, Diffuse unilateral subacute neuroretinitis. 

Morphometric, serologic, and epidemiologic support for 

Baylisascaris as a causative agent: Ophthalmology, v. 100, 

p. 1695–1701.
References Cited 89

Goldberg, N., and Bhatnagar, P., 2012, Letter to the Editor, 

Diffuse unilateral subacute neuroretinitis (DUSN): Ocular 

Immunology & Inflammation, v. 20, p. 139–142.

Goldman, E.A., 1950, Raccoons of North and Middle America: U.S. Fish and Wildlife Service, North American Fauna, 

v. 60, 158 p.

Gozalo, A.S., Cheng, L.I., Maximova, O., St. Claire, M., 

Montali, R.J., Ward, J.M., Elkins, W.R., and Kazacos, 

K.R., 2008, Visceral and neural larva migrans in Rhesus 

macaques: Journal of the American Association for Laboratory Animal Science, v. 47, p. 64–67.

Greve, J.H., 1985, Raccoon ascarids pose public health threat: 

Iowa State University Veterinarian, v. 47, p. 13–14.

Greve, J.H., and O’Brien, S.E., 1989, Adult Baylisascaris

infections in two dogs: Companion Animal Practice, v. 19, 

p. 41–43.

Habermann, R.T., Herman, C.M., and Williams, F.P., Jr., 1958, 

Distemper in raccoons and foxes suspected of having rabies: 

Journal of the American Veterinary Medical Association, 

v. 132, p. 31–35.

Haider, S., Khairnar, K., Martin, D.S., Yang, J., Ralevski, F., 

Kazacos, K.R., and Pillai, D.R., 2012, Possible pet-associated baylisascariasis in child, Canada: Emerging Infectious 

Diseases, v. 18, p. 347–349.

Hajek, J., Yau, Y., Kertes, P., Soman, T., Laughlin, S., Kanani, 

R., Kazacos, K., Dangoudoubiyam, S., and Opavsky, M.A., 

2009, A child with raccoon roundworm meningoencephalitis: A pathogen emerging in your own backyard?: Canadian 

Journal of Infectious Diseases and Medical Microbiology, 

v. 20, p. e177–e180.

Hamann, K.J., Kephart, G.M., Kazacos, K.R., and Gleich, 

G.J., 1989, Immunofluorescent localization of eosinophil 

granule major basic protein in fatal human cases of Baylisascaris procyonis infection: American Journal of Tropical 

Medicine and Hygiene, v. 40, p. 291–297.

Hancock, K., Zajac, A.M., Elvinger, F., and Lindsay, D.S., 

2002, Survey of raccoons (Procyon lotor) in northern Virginia for zoonotic parasites: Proceedings of the American 

Association of Veterinary Parasitologists, v. 47, p. 45.

Hanley, C.S., Simmons, H.A., Wallace, R.S., and Clyde, V.L., 

2006, Visceral and presumptive neural baylisascariasis in an 

orangutan (Pongo pygmaeus): Journal of Zoo and Wildlife 

Medicine, v. 37, p. 553–557.

Harkema, R., and Miller, G.C., 1962, Helminths of Procyon 

lotor solutus from Cape Island, South Carolina: Journal of 

Parasitology, v. 48, p. 333–335.

Harkema, R., and Miller, G.C., 1964, Helminth parasites of the 

raccoon, Procyon lotor, in the southeastern United States: 

Journal of Parasitology, v. 50, p. 60–66.

Helfer, D.H., and Dickinson, E.O., 1976, Parasitic encephalitis 

in pigeons: Avian Diseases, v. 20, p. 209–210.

Herman, C.M., 1939, Parasites obtained from animals in the 

collection of the New York Zoological Park during 1938: 

Zoologica (New York), v. 24, p. 481–485.

Hernandez, S.M., Galbreath, B., Riddle, D.F., Moore, A.P., 

Palamar, M.B., Levy, M.G., DePerno, C.S., Correa, M.T., 

and Yabsley, M.J., 2013, Baylisascaris procyonis in raccoons (Procyon lotor) from North Carolina and current 

status of the parasite in the USA: Parasitology Research, 

v. 112, no. 2, p. 693–698.

Hill, R.E., Jr., Zimmerman, J.J., Greve, J.H., and Beran, G.W., 

1991, Use of ivermectin against several nematodes in naturally infected raccoons (Procyon lotor): Journal of Zoo and 

Wildlife Medicine, v. 22, p. 417–420.

Hirsch, B.T., Prange, S., Hauver, S.A., and Gehrt, S.D., 2014, 

Patterns of latrine use by raccoons (Procyon lotor) and 

implications for Baylisascaris procyonis transmission: Journal of Wildlife Diseases, v. 50, p. 243–249.

Hoberg, E.P., and McGee, S.G., 1982, Helminth parasitism in 

raccoons, Procyon lotor hirtus Nelson and Goldman, in Saskatchewan: Canadian Journal of Zoology, v. 60, p. 53–57.

Hoffman, C.O., and Gottschang, J.L., 1977, Numbers, distribution, and movements of a raccoon population in a suburban residential community: Journal of Mammalogy, 

v. 58, p. 623–636.

Hoffmann, U., Voight, S., and Andreas, U., 2002, Raccoons 

take the offensive. A current assessment, in Kowarik, I., and 

Starfinger, U., eds., Biologische invasionen. Herausforderung zum Handeln?: Neobiota, v. 1, p. 191–192.

Hoffmeister, B., Glaeser, S., Flick, H., Pornschlegel, S., Suttorp, N., and Bergmann, F., 2007, Cerebral toxocariasis after 

consumption of raw duck liver: American Journal of Tropical Medicine and Hygiene, v. 76, p. 600–602.

Huff, D.S., Neafie, R.C., Binder, M.J., De León, G.A., Brown, 

L.W., and Kazacos, K.R., 1984, Case 4. The first fatal Baylisascaris infection in humans: An infant with eosinophilic 

meningoencephalitis: Pediatric Pathology, v. 2, p. 345–352.

Hung, T., Neafie, R.C., and Mackenzie, I.R.A., 2012, 

Baylisascaris procyonis infection in elderly person, British 

Columbia, Canada: Emerging Infectious Diseases, v. 18, 

p. 341–342.

Huntress, S.L., and Spraker, T., 1985, Baylisascaris infection 

in the marmoset: Proceedings of the American Association 

of Zoo Veterinarians, p. 78.
90 Baylisascaris Larva Migrans

Ikeda, T., Asano, M., Matoba, Y., and Abe, G., 2004, Present 

status of invasive alien raccoon and its impact in Japan: 

Global Environmental Research, v. 8, p. 125–131.

Ingle, M.E., Dunbar, S.G., Gathany, M.A., Vasser, M.M., 

Bartsch, J.L., Guffey, K.R., Knox, C.J., Nolan, A.N., Rowlands, C.E., and Trigg, E.C., 2014, Predicting Baylisascaris 

procyonis roundworm prevalence, presence and abundance 

in raccoons (Procyon lotor) of southwestern Ohio using 

landscape features: International Journal for Parasitology: 

Parasites and Wildlife, v. 3, p. 113–117.

Jacobson, H.A., Scanlon, P.F., Nettles, V.F., and Davidson, 

W.R., 1976, Epizootiology of an outbreak of cerebrospinal 

nematodiasis in cottontail rabbits and woodchucks: Journal 

of Wildlife Diseases, v. 12, p. 357–360.

Jacobson, J.E., Kazacos, K.R., and Montague, F.H., Jr., 

1982, Prevalence of eggs of Baylisascaris procyonis

(Nematoda:Ascaroidea) in raccoon scats from an urban and 

a rural community: Journal of Wildlife Diseases, v. 18, 

p. 461–464.

Jardine, C.M., Pearl, D.L., Puskas, K., Campbell, D.G., 

Shirose, L., and Peregrine, A.S., 2014, The impact of land 

use, season, age, and sex on the prevalence and intensity 

of Baylisascaris procyonis infections in raccoons (Procyon 

lotor) from Ontario, Canada: Journal of Wildlife Diseases, 

v. 50, p. 784–791.

Jimenez Martinez, M.A., Cano, E.V., and Rois, J.L., 2015, 

Baylisascaris procyonis larva migrans in two white-headed 

lemurs (Eulemur albifrons) in Spain and response to treatment derived from a human pediatric protocol: Veterinary 

Parasitology, v. 210, p. 246-249. 

Johnson, S.A., 1970, Biology of the raccoon (Procyon lotor 

varius), Nelson and Goldman in Alabama: Auburn, Alabama, Auburn University Agricultural Experiment Station 

Bulletin 402,148 p.

Jones, E.J., and McGinnes, B.S., 1983, Distribution of adult 

Baylisascaris procyonis in raccoons from Virginia: Journal 

of Parasitology, v. 69, p. 653.

Jordan, H.E., and Hayes, F.A., 1959, Gastrointestinal helminths of raccoons (Procyon lotor) from Ossabaw Island, 

Georgia: Journal of Parasitology, v. 45, p. 249–252.

Kahn, L.H., 2006, Confronting zoonoses, linking human and 

veterinary medicine: Emerging Infectious Diseases, v. 12, 

p. 556–561.

Kahn, L.H., Kaplan, B., and Steele, J.H., 2007, Confronting 

zoonoses through closer collaboration between medicine 

and veterinary medicine (as ‘one medicine’): Veterinaria 

Italiana, v. 43, p. 5–19.

Kawanaka, M., Sakamoto, K., and Sugiyama, H., 2001, Raccoon and raccoon roundworm (Baylisascaris procyonis) 

surveillance in Japan: Clinical Parasitology, v. 12, 

p. 121–125. [In Japanese.]

Kawanaka, M., Sakamoto, K., Sugiyama, H., and Morishima, 

Y., 2002, Detection and elimination of Baylisascaris procyonis eggs at zoological gardens exhibiting raccoons in 

Japan, 2000: Infectious Agents Surveillance Report, v. 23, 

p. 202–203. [In Japanese.]

Kazacos, E.A., and Kazacos, K.R., 1988, Pathogenesis and 

pathology of experimental baylisascariasis in ferrets: Proceedings of the American Society of Parasitologists, v. 63, 

p. 60.

Kazacos, K.R., 1981, Animal and public health implications 

of the nematode genus Baylisascaris: Proceedings of the 

American Association of Veterinary Parasitologists, v. 26, 

p. 21.

Kazacos, K.R., 1982, Contaminative ability of Baylisascaris 

procyonis infected raccoons in an outbreak of cerebrospinal 

nematodiasis: Proceedings of the Helminthological Society 

of Washington, v. 49, p. 155–157.

Kazacos, K.R., 1983a, Raccoon roundworms (Baylisascaris 

procyonis). A cause of animal and human disease: West 

Lafayette, In., Purdue University Agricultural Experiment 

Station Bulletin 422, 25 p.

Kazacos, K.R., 1983b, Life cycle studies on Baylisascaris 

procyonis in raccoons: Proceedings of the Conference of 

Research Workers in Animal Diseases, v. 64, p. 24.

Kazacos, K.R., 1983c, Improved method for recovering 

ascarid and other helminth eggs from soil associated with 

epizootics and during survey studies: American Journal of 

Veterinary Research, v. 44, p. 896–900.

Kazacos, K.R., 1984, Baylisascaris infection of human beings: 

West Lafayette, Indiana, Letter to state and territorial public 

health laboratory directors, October 10, 1984.

Kazacos, K.R., 1986, Raccoon ascarids as a cause of larva 

migrans: Parasitology Today, v. 2, p. 253–255.

Kazacos, K.R., 1991, Visceral and ocular larva migrans: Seminars in Veterinary Medicine and Surgery (Small Animal), 

v. 6, p. 227–235.

Kazacos, K.R., 1995, Baylisascariasis, in Farris, R., Mahlow, 

J., Newman, E., Nix, B., eds., Health Hazards in Veterinary 

Practice, 3rd ed.: Austin, Texas Department of Health, 

p. 13–14.
References Cited 91

Kazacos, K.R., 1996a, Baylisascaris procyonis and other helminth larvae as causes of ocular larva migrans and DUSN: 

Proceedings of the International Society of Ophthalmic 

Pathology and the American Association of Ophthalmic 

Pathology, Chicago, p. 20.

Kazacos, K.R., 1996b, Baylisascariasis, in Rudolph, A.M., 

Hoffman, J.I.E., and Rudolph, C.D., eds., Rudolph’s pediatrics, 20th ed.: Stamford, Connecticut, Appleton & Lange, 

p. 716–717.

Kazacos, K.R., 1997, Visceral, ocular, and neural larva 

migrans, in Connor, D.H., Chandler, F.W., Schwartz, D.A., 

Manz, H.J., and Lack, E.E., eds., Pathology of infectious 

diseases, vol. II: Stamford, Connecticut, Appleton & Lange, 

p. 1459–1473.

Kazacos, K.R., 2000, Protecting children from helminthic 

zoonoses: Contemporary Pediatrics, v. 17 (suppl.), p. 1–24.

Kazacos, K.R., 2001, Baylisascaris and related species, in

Samuel, W.M., Pybus, M.J., and Kocan, A.A., eds., Parasitic diseases of wild mammals, 2nd ed.: Ames, Iowa State 

University Press, p. 301–341.

Kazacos, K.R., 2006, Unusual fecal parasite in a dog: 

NAVC Clinician’s Brief, v. 4, p. 37–39.

Kazacos, K.R., Appel, G.O., and Thacker, H.L., 1981, 

Cerebrospinal nematodiasis in a woodchuck suspected of 

having rabies: Journal of the American Veterinary Medical 

Association, v. 179, p. 1102–1104.

Kazacos, K.R., and Boyce, W.M., 1989, Baylisascaris larva 

migrans: Journal of the American Veterinary Medical Association, v. 195, p. 894–903. Addendum, 1995, in Zoonosis 

updates from the Journal of the American Veterinary Medical Association, 2nd ed.: Schaumburg, Illinois, American 

Veterinary Medical Association, p. 29–30.

Kazacos, K.R., Fitzgerald, S.D., and Reed, W.M., 1991, Baylisascaris procyonis as a cause of cerebrospinal nematodiasis 

in ratites: Journal of Zoo and Wildlife Medicine, v. 22, 

p. 460–465.

Kazacos, K.R., Gavin, P.J., Shulman, S.T., Tan, T.Q., Gerber, 

S.I., Kennedy, W.A., Murray, W.J., and Mascola, L., 2002, 

Raccoon roundworm encephalitis—Chicago, Illinois, and 

Los Angeles, California: Morbidity and Mortality Weekly 

Report, v. 50, p. 1153–1155.

Kazacos, K.R., Jelicks, L.A., and Tanowitz, H.B., 2013, Baylisascaris larva migrans, in Garcia, H.H., Tanowitz, H.B., 

and Del Brutto, O.H., eds., Handbook of clinical neurology, 

Vol. 114 (3rd series), Neuroparasitology and tropical neurology: Amsterdam, Elsevier B.V., p. 251–262.

Kazacos, K.R., and Kazacos, E.A., 1984, Experimental infection of domestic swine with Baylisascaris procyonis from 

raccoons: American Journal of Veterinary Research, v. 45, 

p. 1114–1121.

Kazacos, K.R., and Kazacos, E.A., 1988, Diagnostic exercise: 

Neuromuscular condition in rabbits: Laboratory Animal 

Science, v. 38, p. 187–189.

Kazacos, K.R., Kazacos, E.A., Render, J.A., and Thacker, 

H.L., 1982, Cerebrospinal nematodiasis and visceral larva 

migrans in an Australian (Latham’s) brush turkey: Journal 

of the American Veterinary Medical Association, v. 181, 

p. 1295–1298.

Kazacos, K.R., Kazacos, E.A., and Vestre, W.A., 1984, Experimental ocular larva migrans in mice, hamsters, gray squirrels and woodchucks infected with Baylisascaris procyonis: 

Proceedings of the American Association of Veterinary 

Parasitologists, v. 29, p. 27–28.

Kazacos, K.R., Kilbane, T.P., Zimmerman, K.D., Chavez-Lindell, T., Parman, B., Lane, T., Carpenter, L.R., Green, A.L., 

Mann, P.M., Murphy, T.W., Bertucci, B., Gray, A.C., Goldsmith, T.L., Cunningham, M., Stanek, D.R., Blackmore, C., 

Yabsley, M.J., Montgomery, S.P., and Bosserman, E., 2011, 

Raccoon roundworms in pet kinkajous—Three states, 1999 

and 2010: Morbidity and Mortality Weekly Report, v. 60, 

p. 302–305.

Kazacos, K.R., Raymond, L.A., Kazacos, E.A., and Vestre, 

W.A., 1985, The raccoon ascarid. A probable cause of 

human ocular larva migrans: Ophthalmology, v. 92, 

p. 1735–1743.

Kazacos, K.R., Reed, W.M., Kazacos, E.A., and Thacker, 

H.L., 1983, Fatal cerebrospinal disease caused by Baylisascaris procyonis in domestic rabbits: Journal of the American 

Veterinary Medical Association, v. 183, p. 967–971.

Kazacos, K.R., Reed, W.M., and Thacker, H.L., 1986, Cerebrospinal nematodiasis in pheasants: Journal of the American Veterinary Medical Association, v. 189, p. 1353–1354.

Kazacos, K.R., and Turek, J.J., 1983, Scanning electron 

microscopy of the eggs of Baylisascaris procyonis, B. 

transfuga, and Parascaris equorum, and their comparison 

with Toxocara canis and Ascaris suum: Proceedings of the 

Helminthological Society of Washington, v. 50, p. 36–42.

Kazacos, K.R., Vestre, W.A., and Kazacos, E.A., 1984, Raccoon ascarid larvae (Baylisascaris procyonis) as a cause 

of ocular larva migrans: Investigative Ophthalmology and 

Visual Science, v. 25, p. 1177–1183.

Kazacos, K.R., Vestre, W.A., Kazacos, E.A., and Raymond, 

L.A., 1984, Diffuse unilateral subacute neuroretinitis syndrome: Probable cause: Archives of Ophthalmology, v. 102, 

p. 967–968.
92 Baylisascaris Larva Migrans

Kazacos, K.R., Winterfield, R.W., and Thacker, H.L., 1982, 

Etiology and epidemiology of verminous encephalitis in an 

emu: Avian Diseases, v. 26, p. 389–391.

Kazacos, K.R., and Wirtz, W.L., 1983, Experimental cerebrospinal nematodiasis due to Baylisascaris procyonis in 

chickens: Avian Diseases, v. 27, p. 55–65.

Kazacos, K.R., Wirtz, W.L., Burger, P.P., and Christmas, C.S., 

1981, Raccoon ascarid larvae as a cause of fatal central nervous system disease in subhuman primates: Journal of the 

American Veterinary Medical Association, v. 179, 

p. 1089–1094.

Kellner, K.F., Page, L.K., Downey, M., and McCord, S.E., 

2012, Effects of urbanization on prevalence of Baylisascaris procyonis in intermediate host populations: Journal of 

Wildlife Diseases, v. 48, p. 1083–1087.

Kelly, W.R., and Innes, J.R.M., 1966, Cerebrospinal nematodiasis with focal encephalomalacia as a cause of paralysis 

of beavers (Castor canadensis) in the Dublin Zoological 

Gardens: British Veterinary Journal, v. 122, p. 285–287.

Kelly, T.G., Madhavan, V.L., Peters, J.M., Kazacos, K.R., and 

Silvera, V.M., 2012, Spinal cord involvement in a child with 

raccoon roundworm (Baylisascaris procyonis) meningoencephalitis: Pediatric Radiology, v. 42, p. 369–373. 

Kelly, T.G., Silvera, V.M., Madhavan, V.L., Bastos, M.D., 

Kazacos, K.R., Husson, R.N., and Dangoudoubiyam, S., 

2009, Case report: Spinal cord involvement in an infant 

with raccoon roundworm (Baylisascaris procyonis) 

encephalitis: Proceedings of the 47th Annual Meeting of the 

American Society of Neuroradiology, Vancouver British 

Columbia, May 21, 2009, Paper 436.

Kerr, C.L., Henke, S.E., and Pence, D.B., 1997, Baylisascariasis in raccoons from southern coastal Texas: Journal of 

Wildlife Diseases, v. 33, p. 653–655.

Kidder, J.D., 1990, Density and distribution of an urban/suburban raccoon population and the prevalence of patent Baylisascaris procyonis infection: Ithaca, New York, Cornell 

University, M.S. Thesis, 81 p.

Kidder, J.D., Wade, S.E., Richmond, M.E., and Schwager, 

S.J., 1989, Prevalence of patent Baylisascaris procyonis

infection in raccoons (Procyon lotor) in Ithaca, New York: 

Journal of Parasitology, v. 75, p. 870–874.

Kimball, V.P., Young, D., Keith, R., Kazacos, K.R., and 

Sakanari, J., 2003, Occurrence of Baylisascaris procyonis in 

Marin, Sonoma, and Alameda counties in northern California: California Fish and Game, v. 89, p. 46–50.

Koch, F., and Rapp, J., 1981, Zerebrale Nematodiasis bei 

Sumpfbibern (Myocastor coypus)—verursacht durch Larven 

von Baylisascaris (Nematoda): Berliner und Münchener 

Tierärztliche Wochenschrift, v. 94, p. 111–114. [In German.]

Kresta, A.E., Henke, S.E., and Pence, D.B., 2009, Gastrointestinal helminths in raccoons in Texas: Journal of Wildlife 

Diseases, v. 45, p. 1–13.

Kresta, A.E., Henke, S.E., and Pence, D.B., 2010, Baylisascaris procyonis in raccoons in Texas and its relationship to 

habitat characteristics: Journal of Wildlife Diseases, v. 46, 

p. 843–853.

Küchle, M., Knorr, H.L.J., Medenblik-Frysch, S., Weber, A., 

Bauer, C., and Naumann, G.O.H., 1993, Diffuse unilateral 

subacute neuroretinitis syndrome in a German most likely 

caused by the raccoon roundworm, Baylisascaris procyonis: 

Graefe’s Archive for Clinical and Experimental Ophthalmology, v. 231, p. 48–51.

Kwiecien, J.M., Smith, D.A., Key, D.W., Swinton, J., and 

Smith-Maxie, L., 1993, Encephalitis attributed to larval 

migration of Baylisascaris species in emus: Canadian 

Veterinary Journal, v. 34, p. 176–178.

Larson, D.J., and Greve, J.H., 1983, Encephalitis caused by 

Baylisascaris migration in a silver fox: Journal of the American Veterinary Medical Association, v. 183, p. 1274–1275.

Larson, O.R., and Scharf, W.C., 1975, New helminth records 

from Minnesota mammals: Proceedings of the Helminthological Society of Washington, v. 42, p. 174–175.

Lee, A.C.Y., Schantz, P.M., Kazacos, K.R., Montgomery, S.P., 

and Bowman, D.D., 2010, Epidemiologic and zoonotic 

aspects of ascarid infections in dogs and cats: Trends in 

Parasitology, v. 26, p. 155–161.

Leigh, W.H., 1940, Preliminary studies on parasites of upland 

game birds and fur-bearing mammals in Illinois: Bulletin of 

the Illinois Natural History Survey, v. 21, p. 185–194.

Lennox, A.M., Hanika, C., Popielarczyk, M.M., and Kazacos, 

K.R., 2015, Fatal Baylisascaris procyonis infection in a 

household psittacine flock: Proceedings of the Association 

of Exotic Mammal Veterinarians Annual Conference, San 

Antonio, Texas, v. 14, Session #114.

Lewis, R.A., 1985, Discussion (of The raccoon ascarid. A 

probable cause of human ocular larva migrans): Ophthalmology, v. 92, p. 1743–1744.

Lindquist, W.D., 1978, Baylisascaris procyonis for testing 

anthelmintics against migratory ascarids: American Journal 

of Veterinary Research, v. 39, p. 1868–1869.
References Cited 93

Liu, G., Fennelly, G., Kazacos, K.R., Grose, C., Dobroszycki, 

J., Saffra, N., Coyle, C.M., Weiss, L.M., Szlechter, M.M., 

and Tanowitz, H.B., 2015, Case report: Baylisascaris 

procyonis and Herpes simplex Virus 2 co-infection presenting as ocular larva migrans with granuloma formation in a 

child: American Journal of Tropical Medicine and Hygiene, 

v. 93, p. 612–614.

LoGiudice, K., 1995, Control of Baylisascaris procyonis

(Nematoda) in raccoons (Procyon lotor) through the use of 

anthelmintic baits: A potential method for reducing mortality in the Allegheny woodrat (Neotoma floridana magister): 

New Brunswick, New Jersey, Rutgers University, M.S. 

Thesis, 78 p.

LoGiudice, K., 2001, Latrine foraging strategies of two small 

mammals: Implications for the transmission of Baylisascaris procyonis: American Midland Naturalist, v. 146, 

p. 369–378.

LoGiudice, K., 2003, Trophically transmitted parasites and the 

conservation of small populations: Raccoon roundworm and 

the imperiled Allegheny woodrat: Conservation Biology, 

v. 17, p. 258–266.

LoGiudice, K., 2006, Toward a synthetic view of extinction: A 

history lesson from a North American rodent: BioScience, 

v. 56, p. 687–693.

Long, D.B., Campbell, T.A., and Henke, S.E., 2006, Baylisascaris procyonis (Nematoda: Ascaridoidea) in raccoons 

(Procyon lotor) from Duval County, Texas: Texas Journal of 

Science, v. 58, p. 281–285.

Loretti, A.P., Dangoudoubiyam, S., Koobs, J., Gai, J.J., and 

Kazacos, K.R., 2008, Cases of cerebral nematodiasis in 

canaries and emus due to Baylisascaris procyonis larval 

migration (neural larva migrans) diagnosed in the State of 

California, year 2007: Proceedings of the American Association of Veterinary Laboratory Diagnosticians, v. 51, 

p. 103.

Lotze, J-H., and Anderson, S., 1979, Procyon lotor: Mammalian Species, v. 119, p. 1–8. 

Lutz, W., 1984, Die Verbreitung des Waschbären im mittleuropäischen Raum: Zeitshcrift fur Jagdwissenschaft, v. 30, 

p. 218–228. [In German.]

Lutz, W., 1996, The introduced raccoon Procyon lotor population in Germany: Wildlife Biology, v. 2, p. 228.

Lux, E., and Priemer, J., 1995, Zur Parasitierung wildlebender 

Waschbären unter dem Aspekt ihrer nordamerikanischen: 

Verhandlungsbericht Erkrankungen der Zootiere, v. 37, 

p. 429–434. [In German.]

Lyons, E.T., Swerczek, T.W., and Tolliver, S.C., 2001, Central 

nervous system disorder in a woodchuck (Marmota monax) 

in central Kentucky caused by a nematode in the genus 

Baylisascaris: Journal of the Kentucky Academy of 

Science, v. 62, p. 142–144.

Mackay, A., Robitaille, J., Messier, S., and Villeneuve, A., 

1995, Baylisascaris chez le raton laveur au Québec: possibilité de zoonose: Le Médecin Vétérinaire du Québec, 

v. 25, p. 102–105. [In French.]

Matoba, Y., Yamada, D., Asano, M., Oku, Y., Kitaura, K., 

Yagi, K., Tenora, F., and Asakawa, M., 2006, Parasitic 

helminths from feral raccoons (Procyon lotor) in Japan: 

Helminthologia, v. 43, p. 139–146.

Matoff, K., and Komandarev, S., 1965, Comparative studies 

on the migration of the larvae of Toxascaris leonina and 

Toxascaris transfuga: Zeitschrift fur Parasitenkunde, v. 25, 

p. 538–555.

McCleery, R.A., Foster, G.W., Lopez, R.R., Peterson, M.J., 

Forrester, D.J., and Silvy, N.J., 2005, Survey of raccoons 

on Key Largo, Florida, USA, for Baylisascaris procyonis: 

Journal of Wildlife Diseases, v. 41, p. 250–252.

McClure, G.W., 1933, Nematode parasites of mammals. From 

specimens collected in the New York Zoological Park, 

1931: Zoologica (New York), v. 15, p. 29–47.

McFadden, K.W., Wade, S.E., Dubovi, E.J., and Gompper, 

M.E., 2005, A serological and fecal parasitologic survey of 

the critically endangered pygmy raccoon (Procyon pygmaeus): Journal of Wildlife Diseases, v. 41, p. 615–617.

McGowan, E.M., 1993, Experimental release and fate study of 

the Allegheny woodrat (Neotoma magister): Albany, New 

York Federal Aid Project W–166–E; E–1, Job No. VIII–7, 

New York State Department of Environmental Conservation, 15 p.

McNeil, C.W., and Krogsdale, J.T., 1953, Parasites of raccoons in southwest Washington: Journal of Mammalogy, 

v. 34, p. 123–124.

Medway, W., Skand, D.L., and Sarver, C.F., 1989, Neurologic 

signs in American porcupines (Erethizon dorsatum) infected 

with Baylisascaris and Toxoplasma: Journal of Zoo and 

Wildlife Medicine, v. 20, p. 207–211.

Mehta, P., Boyd, Z., and Cully, B., 2010, Raccoon roundworm 

encephalitis: Pediatric Radiology, v. 40, p. 1834–1836.

Mets, M.B., Noble, A.G., Basti, S., Gavin, P., Davis, A.T., 

Shulman, S.T., and Kazacos, K.R., 2003, Eye findings of 

diffuse unilateral subacute neuroretinitis and multiple choroidal infiltrates associated with neural larva migrans due to 

Baylisascaris procyonis: American Journal of Ophthalmology, v. 135, p. 888–890.
94 Baylisascaris Larva Migrans

Mitchell, M.A., Hungerford, L.L., Nixon, C., Sullivan, J., 

Esker, T., and Koerkenmeier, R., 1998, Prevalence patterns 

of patent Baylisascaris procyonis infections in raccoons 

(Procyon lotor) in west-central Illinois: Proceedings of the 

American Association of Zoo Veterinarians and American 

Association of Wildlife Veterinarians Joint Conference, 

p. 230–233.

Miyashita, M., 1993, Prevalence of Baylisascaris procyonis in 

raccoons in Japan and experimental infections of the worm 

to laboratory animals: Seikatsu Eisei (Journal of Urban Living and Health Association), v. 37, p. 137–151. [In Japanese 

with English summary.]

Moertel, C.L., Kazacos, K.R., Butterfield, J.H., Kita, H., Watterson, J., and Gleich, G.J., 2001, Eosinophil-associated 

inflammation and elaboration of eosinophil-derived proteins 

in 2 children with raccoon roundworm (Baylisascaris procyonis) encephalitis: Pediatrics, v. 108, p. e93–e99, accessed 

July 30, 2014, at http://www.pediatrics.org/cgi/content/

full/108/5/e93. 

Moore, B.E., 2009, A case of cerebral baylisascariasis: Neuropathology Blog, September 28, 2009, accessed July 30, 

2014, at http://neuropathologyblog.blogspot.com/2009/09/

case-of-cerebral-baylisascariasis.html.

Moore, L., Ash, L., Sorvillo, F., and Berlin, O.G.W., 2004, 

Baylisascaris procyonis in California: Emerging Infectious 

Diseases, v. 10, p. 1693–1694.

Morgan, B.B., and Waller, E.F., 1940, Severe parasitism in a 

raccoon, (Procyon lotor lotor Linnaeus): Transactions of the 

American Microscopical Society, v. 59, p. 523–527.

Müller-Using, D., 1959, Die Ausbreitung des Waschbären 

in Westdeutschland: Z. Jagdwiss, v. 5, p. 108–109. 

[In German.]

Murray, M., Nelson, G., and Smith, M., 2005, Raccoon roundworm (Baylisascaris procyonis) survey: Reno, Nevada, 

Washoe County District Health Department Vector-Borne 

Diseases and Nevada Department of Agriculture Animal 

Disease and Food Safety Laboratory, accessed September 

10, 2014, at https://www.washoecounty.us/health/programsand-services/vector-borne-diseases/surveillance_activities/

raccoon-roundworm-survey.php

Murray, W.J., 2004, Raccoon roundworm infection (Baylisascariasis): A zoonosis of pediatric and public health concern, 

in Scheld, W.M., Murray, B.E., and Hughes, J.M., eds., 

Emerging Infections 6: Washington, D.C., ASM Press, 

p. 159–175.

Murray, W.J., and Kazacos, K.R., 2004, Raccoon roundworm 

encephalitis: Clinical Infectious Diseases, v. 39, 

p. 1484–1492.

Myers, R.K., Monroe, W.E., and Greve, J.H., 1983, Cerebrospinal nematodiasis in a cockatiel: Journal of the American 

Veterinary Medical Association, v. 183, p. 1089–1090.

Nadler, S.A., and Hudspeth, D.S.S., 2000, Phylogeny of the 

Ascaridoidea (Nematoda: Ascaridida) based on three genes 

and morphology: Hypotheses of structural and sequence 

evolution: Journal of Parasitology, v. 86, p. 380–393.

Nadler, S., Bieberich, A., Pagan, C., Naem, S., and Richardson, D., 2010, Evolutionary genetics and geographic variation of Baylisascaris species in North America: Proceedings 

of the American Society of Parasitologists, v. 85, p. 84.

Nelson, G.S., 1983, Teaching medical parasitology, in Warren, 

K.S. and Bowers, J.Z., eds., Parasitology—a global perspective: New York, New York, Springer-Verlag, p. 158–168.

Nettles, V.F., Davidson, W.R., Fisk, S.K., and Jacobson, H.A., 

1975, An epizootic of cerebrospinal nematodiasis in cottontail rabbits: Journal of the American Veterinary Medical 

Association, v. 167, p. 600–602.

Nichols, R.L., 1956, The etiology of visceral larva migrans. I. 

Diagnostic morphology of infective second-stage Toxocara

larvae: Journal of Parasitology, v. 42, p. 349–362.

Noce, S.M., Ayebo, A., Kazacos, K., Humphreys, J., and 

Brenneman, W.M., 2007, Prevalence and intensity of Baylisascaris procyonis (raccoon roundworm) in Indiana county, 

Pennsylvania: Journal of the Pennsylvania Academy of 

Science, v. 80, p. 57–60.

Nupp, T.E., and Swihart, R.K., 2000, Landscape-level correlates of small-mammal assemblages in forest fragments of 

farmland: Journal of Mammalogy, v. 81, p. 512–526.

Ochiai, K., Ishii, M., and Furukawa, T., 2002, Invasion and 

distribution of the raccoon, Procyon lotor, in Chiba Prefecture, central Japan: Journal of the Natural History Museum 

and Institute, Chiba, v. 7, p. 21–27. [In Japanese.]

Olsen, O.W., and Fenstermacher, R., 1938, The raccoon, a new host of Ascaris columnaris Leidy, 1856 

(Nematoda:Ascaridae): Proceedings of the Helminthological Society of Washington, v. 5, p. 20.

Overstreet, R.M., 1970, Baylisascaris procyonis (Stefanski 

and Zarnowski, 1951) from the kinkajou, Potos flavus, in 

Colombia: Proceedings of the Helminthological Society of 

Washington, v. 37, p. 192–195.

Owen, S.F., Edwards, J.W., Ford, W.M., Crum, J.M., and 

Wood, P.B., 2004, Raccoon roundworm in raccoons in 

central West Virginia: Northeastern Naturalist, v. 11, 

p. 137–142.

Page, L.K., 1998, Ecology and transmission dynamics of 

Baylisascaris procyonis: West Lafayette, Indiana, Purdue 

University, Ph.D. Thesis, 138 p.
References Cited 95

Page, L.K., 2013, Parasites and the conservation of small 

populations: The case of Baylisascaris procyonis: International Journal for Parasitology: Parasites and Wildlife, v. 2, 

p. 203–210.

Page, L.K., Anchor, C., Luy, E., Kron, S., Larson, G., Madsen, 

L., Kellner, K., and Smyser, T.J., 2009, Backyard raccoon 

latrines and risk for Baylisascaris procyonis transmission 

to humans: Emerging Infectious Diseases, v. 15, 

p. 1530–1531.

Page, L.K., Beasley, J.C., Olson, Z.H., Smyser, T.J., Downey, 

M., Kellner, K.F., McCord, S.E., Egan, T.S., II, and Rhodes, 

O.E., Jr., 2011, Reducing Baylisascaris procyonis roundworm larvae in raccoon latrines: Emerging Infectious 

Diseases, v. 17, p. 90–93.

Page, L.K., Delzell, D.A.P., Gehrt, S.D., Dunkerton, E., Hiben, 

M., Walter, E., and Kazacos, K.R., 2016, The structure and 

seasonality of Baylisascaris procyonis populations in raccoons: Journal of Wildlife Diseases, v. 52, at http://www.

jwildlifedis.org/doi/pdf/10.7589/2015-06-153.

Page, L.K., Gehrt, S.D., Cascione, A., and Kellner, K.F., 2009, 

The relationship between Baylisascaris procyonis prevalence and raccoon population structure: Journal of Parasitology, v. 95, p. 1314–1320.

Page, L.K., Gehrt, S.D., and Robinson, N.P., 2008, Land-use 

effects on prevalence of raccoon roundworm (Baylisascaris 

procyonis): Journal of Wildlife Diseases, v. 44, p. 594–599.

Page, L.K., Gehrt, S.D., Titcombe, K.K., and Robinson, N.P., 

2005, Measuring prevalence of raccoon roundworm (Baylisascaris procyonis): A comparison of common techniques: 

Wildlife Society Bulletin, v. 33, p. 1406–1412.

Page, L.K., Johnson, S.A., Swihart, R.K., and Kazacos, K.R., 

2012, Prevalence of Baylisascaris procyonis in habitat 

associated with Allegheny woodrat (Neotoma magister) 

populations in Indiana: Journal of Wildlife Diseases, v. 48, 

p. 503–507.

Page, L.K., Swihart, R.K., and Kazacos, K.R., 1998, Raccoon 

latrine structure and its potential role in transmission of 

Baylisascaris procyonis to vertebrates: American Midland 

Naturalist, v. 140, p. 180–185.

Page, L.K., Swihart, R.K., and Kazacos, K.R., 1999, Implications of raccoon latrines in the epizootiology of baylisascariasis: Journal of Wildlife Diseases, v. 35, p. 474–480.

Page, L.K., Swihart, R.K., and Kazacos, K.R., 2001a, Seed 

preferences and foraging by granivores at raccoon latrines 

in the transmission dynamics of the raccoon roundworm 

(Baylisascaris procyonis): Canadian Journal of Zoology, 

v. 79, p. 616–622.

Page, L.K., Swihart, R.K., and Kazacos, K.R., 2001b, Foraging among feces: Food availability affects parasitism of 

Peromyscus leucopus by Baylisascaris procyonis: Journal 

of Mammalogy, v. 82, p. 993–1002.

Page, L.K., Swihart, and R.K., Kazacos, K.R., 2001c, Changes 

in transmission of Baylisascaris procyonis to intermediate 

hosts as a function of spatial scale: Oikos, v. 93, p. 213–220.

Pai, P.J., Blackburn, B.G., Kazacos, K.R., Warrier, R.P. and 

Bégué, R.E., 2007, Full recovery from Baylisascaris procyonis eosinophilic meningitis: Emerging Infectious Diseases, 

v. 13, p. 928–930.

Papini, R., and Casarosa, L., 1994, A report on the pathology 

of Baylisascaris transfuga (Ascarididae, Nematoda) for 

mice: Revue de Médecine Vétérinaire, v. 145, p. 949–952.

Papini, R., Cavicchio, P., and Casarosa, L., 1993, Experimental infection in chickens with larvae of Baylisascaris 

transfuga (Nematoda: Ascaridoidea): Folia Parasitologica, 

v. 40, p. 141–143.

Papini, R., Demi, S., and Della Croce, G., 1996, Observations 

on the migratory behaviour of Baylisascaris transfuga

larvae in rabbits: Revue de Médecine Vétérinaire, v. 147, 

p. 893–896. 

Papini, R., Renzoni, G., Lo Piccolo, S., and Casarosa, L., 

1996, Ocular larva migrans and histopathological lesions in 

mice experimentally infected with Baylisascaris transfuga

embryonated eggs: Veterinary Parasitology, v. 61, 

p. 315–320.

Papini, R., Renzoni, G., Malloggi, M., and Casarosa, L., 1994, 

Visceral larva migrans in mice experimentally infected 

with Baylisascaris transfuga (Ascarididae: Nematoda): 

Parasitologia, v. 36, p. 321–329.

Park, C., Levee, D.J., Gilbreath, S., Parman, R., Frazer, L., 

Kaufman, R., and Murray, W.J., 1998, A survey of the 

prevalence of the raccoon ascarid, Baylisascaris procyonis, 

in a geographically defined population of suburban and 

urban raccoons: Proceedings of the American Society for 

Microbiology, v. 98, p. 547.

Park, S.Y., Glaser, C., Murray, W.J., Kazacos, K.R., Rowley, 

H.A., Fredrick, D.R., and Bass, N., 2000, Raccoon roundworm (Baylisascaris procyonis) encephalitis: Case report 

and field investigation: Pediatrics, v. 106, p. e56–e60, 

accessed July 30, 2014, at http://www.pediatrics.org/cgi/

content/full/106/4/e56.

Parkanzky, M.C., 2015, Baylisascaris spp. in non-raccoon 

procyonid hosts and assessment of potential risk of human 

exposure: West Lafayette, Indiana, Purdue University, M.S. 

Thesis, 61 p.
96 Baylisascaris Larva Migrans

Parsons, H.E., 1952, Nematode chorioretinitis. Report of a 

case, with photographs of a viable worm: Archives of Ophthalmology, v. 47, p. 799–800.

Pegg, E.J., 1977, A new approach to the control of Toxocara 

canis and other parasitic ova on concrete-floored kennel 

runs: British Veterinary Journal, v. 133, p. 427–431.

Perlman, J.E., Kazacos, K.R., Imperato, G.H., Desai, R.U., 

Schulman, S.K., Edwards, J., Pontrelli, L.R., Machado, F.S., 

Tanowitz, H.B., and Saffra, N.A., 2010, Baylisascaris procyonis neural larva migrans in an infant in New York City: 

Journal of Neuroparasitology, v. 1, 5 p.

Pessier, A.P., Stringfield, C., Tragle, J., Holshuh, H.J., Nichols, 

D.K., and Montali, R.J., 1997, Cerebrospinal nematodiasis 

due to Baylisascaris sp. in golden-headed lion tamarins 

(Leontopithecus chrysomelas): implications for management: Proceedings of the American Association of Zoo 

Veterinarians, p. 245–247.

Peters, J.M., Madhavan, V.L., Kazacos, K.R., Husson, R.N., 

Dangoudoubiyam, S., and Soul, J.S., 2012, Good outcome 

with early empiric treatment of neural larva migrans due to 

Baylisascaris procyonis: Pediatrics, v. 129, p. e806–e811, 

accessed July 30, 2014, at http://www.pediatrics.org/cgi/

content/full/129/3/e806.

Peterson, R.T., 1980, A field guide to the birds (4th ed.): New 

York, Houghton Mifflin Co., 384 p.

Peterson, R.T., 1990, A field guide to western birds (3rd ed.): 

New York, Houghton Mifflin Co., 432 p.

Pigage, J.C., Pigage, H.K., and Todd, K.S., Jr., 1983, Intestinal 

parasites of mammals from Willowbrook Wildlife Haven: A 

survey with implications for management: Proceedings of 

the National Wildlife Rehabilitation Association, v. 2, 

p. 174–184.

Pipas, M.J., Page, L.K., and Kazacos, K.R., 2014, Surveillance 

for Baylisascaris procyonis in raccoons (Procyon lotor) 

from Wyoming, USA: Journal of Wildlife Diseases, v. 50, 

p. 777–783.

Popiotek, M., Szczęsna-Staśkiewicz, J., Bartoszewicz, M., 

Okarma, H., Smalec, B., and Zalewski, A., 2011, Helminth 

parasites of an introduced invasive carnivore species, the 

raccoon (Procyon lotor L.), from the Warta Mouth National 

Park (Poland): Journal of Parasitology, v. 97, p. 357–360.

Prange, S., Gehrt, S.D., and Wiggers, E.P., 2003, Demographic 

factors contributing to high raccoon densities in urban landscapes: Journal of Wildlife Management, v. 67, p. 324–333.

Prange, S., Gehrt, S.D., and Wiggers, E.P., 2004, Influences of 

anthropogenic resources on raccoon movements and spatial 

distribution in urbanized systems: Journal of Mammalogy, 

v. 85, p. 483–490.

Price, R.L., and Harman, D.M., 1983, Helminths from the raccoon, Procyon lotor litoreus Nelson and Goldman 1930, on 

St. Catherines Island, Georgia: Proceedings of the Helminthological Society of Washington, v. 50, p. 343–344.

Rascoe, L.N., Santamaria, C., Handali, S., Dangoudoubiyam, S., Kazacos, K.R., Wilkins, P.P., and Ndao, M., 2013, 

Interlaboratory optimization and evaluation of a serological 

assay for diagnosis of human baylisascariasis: Clinical and 

Vaccine Immunology, v. 20, p. 1758–1763.

Rausch, R., 1946, The raccoon, a new host for Microphallus

sp., with additional notes on M. ovatus from turtles: Journal 

of Parasitology, v. 32, p. 208–209.

Raymond, L.A., Gutierrez, Y., Strong, L.E., Wander, A.H., 

Buten, R., and Cordan, D., 1978, Living retinal nematode 

(filarial-like) destroyed with photocoagulation: Ophthalmology, v. 85, p. 944–949.

Reed, C., Henke, S.E., and Kresta, A.E., 2012, Frequency of 

deposition and location of Baylisascaris procyonis eggs 

in raccoon feces: Journal of Wildlife Diseases, v. 48, 

p. 190–194.

Reed, W.M., Kazacos, K.R., Dhillon, A.S., Winterfield, R.W., 

and Thacker, H.L., 1981, Cerebrospinal nematodiasis in 

bobwhite quail: Avian Diseases, v. 25, p. 1039–1046.

Reilly, E., 2008, ‘No family deserves this’. Raccoon roundworm robs toddler of sight, mobility: The Hamilton 

Spectator, Saturday, October 4, 2008, A16.

Reinemeyer, C.R., Berthelin-Baker, C., Alva, R., Erwin, J., 

Larsen, D., Romano, D., Pettyjohn, M., Ulrich, M.E., Manthei, D., and Kazacos, K.R., 2008, A controlled evaluation 

of the efficacy of HEARTGARD PLUS® against induced 

Baylisascaris procyonis infection in dogs: Proceedings of 

the American Association of Veterinary Parasitologists, 

v. 53, p. 36.

Richardson, D.J., Owen, W.B., and Snyder, D.E., 1992, Helminth parasites of the raccoon (Procyon lotor) from northcentral Arkansas: Journal of Parasitology, v. 78, p. 163–166.

Richardson, J.A., Kazacos, K.R., Thacker, H.L., Dhillon, A.S., 

and Winterfield, R.W., 1980, Verminous encephalitis in 

commercial chickens: Avian Diseases, v. 24, p. 498–503.

Richter, C.B., and Kradel, D.C., 1964, Cerebrospinal 

nematodosis in Pennsylvania groundhogs (Marmota 

monax): American Journal of Veterinary Research, v. 25, 

p. 1230–1235.

Riley, S.P.D., Hadidian, J., and Manski, D.A., 1998, 

Population density, survival, and rabies in raccoons in an 

urban national park: Canadian Journal of Zoology, v. 76, 

p. 1153–1164.
References Cited 97

Robel, R.J., Barnes, N.A., and Upton, S.J., 1989, Gastrointestinal helminths and protozoa from two raccoon populations 

in Kansas: Journal of Parasitology, v. 75, p. 1000–1003.

Robinson, V.B., Newberne, J.W., and Brooks, D.M., 1957, 

Distemper in the American raccoon (Procyon lotor): Journal 

of the American Veterinary Medical Association, v. 131, 

p. 276–278.

Rosatte, R.C., Powers, M.J., and MacInnes, C.D., 1991, Ecology of urban skunks, raccoons and foxes in metropolitan 

Toronto, in Adams, L.W., and Leedy, D.L., eds., Wildlife 

conservation in metropolitan environments: Columbia, 

Maryland, National Institute of Urban Wildlife, p. 31–38.

Roth, L., Georgi, M.E., King, J.M., and Tennant, B.C., 1982, 

Parasitic encephalitis due to Baylisascaris sp. in wild and 

captive woodchucks (Marmota monax): Veterinary Pathology, v. 19, p. 658–662.

Roussere, G.P., Murray, W.J., Raudenbush, C.B., Kutilek, 

M.J., Levee, D.J., and Kazacos, K.R., 2003, Raccoon roundworm eggs near homes and risk for larva migrans disease, 

California communities: Emerging Infectious Diseases, v. 9, 

p. 1516–1522.

Rowley, H.A., Uht, R.M., Kazacos, K.R., Sakanari, J., 

Wheaton, W.V., Barkovich, A.J., and Bollen, A.W., 2000, 

Radiologic-pathologic findings in raccoon roundworm 

(Baylisascaris procyonis) encephalitis: American Journal of 

Neuroradiology, v. 21, p. 415–420.

Rudmann, D.G., Kazacos, K.R., Storandt, S.T., Harris, D.L., 

and Janovitz, E.B., 1996, Baylisascaris procyonis larva 

migrans in a puppy: A case report and update for the veterinarian: Journal of the American Animal Hospital Association, v. 32, p. 73–76.

Russell, D.J., 2006, Avian neural larva migrans due to Baylisascaris procyonis: Natural and experimental infection in 

psittacine birds: Guelph, Ontario, University of Guelph, 

DVSc Thesis, 284 p.

Russell, D.J., Smith, D.A., Crawshaw, G., Dutton, C.J., ArneOlberg, R., Bertelsen, M.F., and Rose, K., 2005, Neural 

larva migrans due to Baylisascaris procyonis in captive 

lovebirds and lorikeets: A case series: Proceedings of the 

Association of Zoo Veterinarians, p. 135.

Saffra, N.A., Perlman, J.E., Desai, R.U., Kazacos, K.R., 

Coyle, C.M., Machado, F.S., Kedhar, S.R., Engelbert, M., 

and Tanowitz, H.B., 2010, Baylisascaris procyonis induced 

diffuse unilateral subacute neuroretinitis in New York City: 

Journal of Neuroparasitology, v. 1, 4 p. 

Sakla, A.A., Donnelly, J.J., Khatami, M., and Rockey, J.H., 

1989, Baylisascaris procyonis (Stefanski and Zarnowski, 

1951) Ascarididae: Nematoda. I. Embryonic development 

and morphogenesis of second stage larvae: Assiut Veterinary Medical Journal, v. 21, p. 68–76.

Samson, A., Dubay, S.A., Huspeni, T.C., and Cyr, A., 2012, 

Influence of environmental variables on Baylisascaris procyonis infection in raccoons: Journal of Parasitology, v. 98, 

p. 1279–1282.

Sanders, M.P., 2015, The worm conqueror: Veterinary 

Heritage, v. 38, p.36–45.

Sanford, S.E., 1991, Cerebrospinal nematodiasis caused by 

Baylisascaris procyonis in chinchillas: Journal of Veterinary 

Diagnostic Investigation, v. 3, p. 77–79.

Sass, B., and Gorgacz, E.J., 1978, Cerebral nematodiasis in a 

chukar partridge: Journal of the American Veterinary Medical Association, v. 173, p. 1248–1249.

Sato, H., Furuoka, H., and Kamiya, H., 2002, First outbreak of 

Baylisascaris procyonis larva migrans in rabbits in Japan: 

Parasitology International, v. 51, p. 105–108.

Sato, H., Kamiya, H., and Furuoka, H., 2003, Epidemiological 

aspects of the first outbreak of Baylisascaris procyonis larva 

migrans in rabbits in Japan: Journal of Veterinary Medical 

Science, v. 65, p. 453–457.

Sato, H., Matsuo, K., Osanai, A., Kamiya, H., Akao, N., 

Owaki, S., and Furuoka, H., 2004, Larva migrans by 

Baylisascaris transfuga: Fatal neurological diseases in 

Mongolian jirds, but not mice: Journal of Parasitology, 

v. 90, p. 774–781.

Sato, H., and Suzuki, K., 2006, Gastrointestinal helminths of 

feral raccoons (Procyon lotor) in Wakayama Prefecture, 

Japan: Journal of Veterinary Medical Science, v. 68, 

p. 311–318.

Sato, H., Une, Y., Kawakami, S., Saito, E., Kamiya, H., 

Akao, N., and Furuoka, H., 2005, Fatal Baylisascaris larva 

migrans in a colony of Japanese macaques kept by a 

safari-style zoo in Japan: Journal of Parasitology, v. 91, 

p. 716–719.

Schaffer, G.D., Davidson, W.R., Nettles, V.F., and Rollor, 

E.A., III, 1981, Helminth parasites of translocated raccoons 

(Procyon lotor) in the southeastern United States: Journal of 

Wildlife Diseases, v. 17, p. 217–227.

Schrott, E.R., 1961, Parasitäre Netzhauterkrankung durch 

Askaris: Verhandlungen der Österreichischen Ophthalmologischen Gesellschaft, v. 6, p. 160–165. [In German.]
98 Baylisascaris Larva Migrans

Schueler, R.L., 1973, Cerebral nematodiasis in a red squirrel: 

Journal of Wildlife Diseases, v. 9, p. 58–60.

Schultz, A.L., 1962, A survey of parasites of the raccoon 

(Procyon lotor) in southeastern Michigan: Ann Arbor, The 

University of Michigan, M.S. Thesis, 42 p.

Schultz, T., 2002, ‘Raccoon roundworm’ infection confirmed: 

Santa Barbara, CA: Santa Barbara News-Press, 

May 8, 2002.

Senger, C.M., and Neiland, K.A., 1955, Helminth parasites of 

some fur-bearers of Oregon: Journal of Parasitology, v. 41, 

p. 637–638.

Sexsmith, J.L., Whiting, T.L., Green, C., Orvis, S., Berezanski, 

D.J., and Thompson, A.B., 2009, Prevalence and distribution of Baylisascaris procyonis in urban raccoons (Procyon 

lotor) in Winnipeg, Manitoba: Canadian Veterinary Journal, 

v. 50, p. 846–850.

Shafir, S.C., Sorvillo, F.J., Sorvillo, T., and Eberhard, M.L., 

2011, Viability of Baylisascaris procyonis eggs: Emerging 

Infectious Diseases, v. 17, p. 1293–1295.

Shafir, S.C., Wang, W., Sorvillo, F.J., Wise, M.E., Moore, 

L., Sorvillo, T., and Eberhard, M.L., 2007, Thermal death 

point of Baylisascaris procyonis eggs: Emerging Infectious 

Diseases, v. 13, p. 172–173.

Shafir, S.C., Wise, M.E., Sorvillo, F.J., and Ash, L.R., 2006, 

Central nervous system and eye manifestations of infection 

with Baylisascaris procyonis: Current Infectious Disease 

Reports, v. 8, p. 307–313.

Sheppard, C.H., 1995, Susceptibility of Peromyscus leucopus

(white-footed mouse) and Mus musculus (albino mouse) 

to infection with Baylisascaris procyonis: West Lafayette, 

Indiana, Purdue University, M.S. Thesis, 108 p.

Sheppard, C.H., 1996, Verhalten und Pathogenität der Larven 

von Baylisascaris procyonis Stefanski und Zarnowski 1951 

(Ascarididae) in verschiedenen Mäusearten: Hannover, 

Germany, Tierärztliche Hochschule Hannover, Dissertation, 

Doctor Medicinae Veterinariae, 100 p. [In German.]

Sheppard, C.H., and Kazacos, K.R., 1997, Susceptibility of 

Peromyscus leucopus and Mus musculus to infection with 

Baylisascaris procyonis: Journal of Parasitology, v. 83, 

p. 1104–1111.

Shoieb, A., and Radi, Z.A., 2014, Cerebral Baylisascaris larva 

migrans in a cynomolgus macaque (Macaca fascicularis): 

Experimental and Toxicologic Pathology, v. 66, p. 263–265.

Singaravelu, K., Coyle, C.M., Martins, Y.C., Weiss, L.M., 

Machado, F.S., Tanowitz, H.B., and Kazacos, K.R., 2016, 

Nematode larva migrans, in Singh, S.K., ed., Human emerging and re-emerging infections (1st ed.): Viral & parasitic 

infections: Hoboken, New Jersey, John Wiley & Sons, v. 1, 

p. 453–480.

Sivalingam, A., Goldberg, R.E., Augsburger, J., and Frank, P., 

1991, Diffuse unilateral subacute neuroretinitis: Archives 

of Ophthalmology, v. 109, p. 1028.

Smiley, D., 1977a, Research report—Raccoon: New Paltz, 

New York, Mohonk Preserve, Daniel Smiley Research 

Center, 19 p.

Smiley, D., 1977b, Research report—Cave (Wood) Rat: 

New Paltz, New York, Mohonk Preserve, Daniel Smiley 

Research Center, 5 p.

Smiley, D., and Huth, P.C., 1986, “Crazy” animals: New Paltz, 

New York, Mohonk Preserve, Daniel Smiley Research 

Center, Natural Science Note 115, 2 p.

Smith, D.A., Kwiecien, J.M., and Smith-Maxie, L., 1993, 

Encephalitis in emus resulting from migration of Baylisascaris sp: Proceedings of the Association of Avian 

Veterinarians, p. 301–303.

Smith, H.T., and Engeman, R.M., 2002, An extraordinary 

raccoon, Procyon lotor, density at an urban park: Canadian 

Field-Naturalist, v. 116, p. 636–639.

Smith, R.A., Kennedy, M.L., and Wilhelm, W.E., 1985, 

Helminth parasites of the raccoon (Procyon lotor) from 

Tennessee and Kentucky: Journal of Parasitology, v. 71, 

p. 599–603.

Smith, S.L., 1992, The presence of the raccoon roundworm, 

Baylisascaris procyonis in Procyon lotor in Nova Scotia: 

Wolfville, Nova Scotia, Acadia University, Senior Honors 

Thesis, 12 p.

Smyser, T.J., Johnson, S.A., Page, L.K., Hudson, C.M., and 

Rhodes, O.E., Jr., 2013, Use of experimental translocations 

of Allegheny woodrat to decipher causal agents of decline: 

Conservation Biology, v. 27, p. 752–762.

Smyser, T.J., Johnson, S.R., Stallard, M.D., McGrew, A.K., 

Page, L.K., Crider, N., Ballweber, L.R., Swihart, R.K., 

and VerCauteren, K.C., 2015, Evaluation of anthelmintic 

fishmeal polymer baits for the control of Baylisascaris 

procyonis in free-ranging raccoons (Procyon lotor): Journal 

of Wildlife Diseases, v. 51, p. 640–650.
References Cited 99

Smyser, T.J., Page, L.K., Johnson, S.A., Hudson, C.M., Kellner, K.F., Swihart, R.K., and Rhodes, O.E., Jr., 2013, Management of raccoon roundworm in free-ranging raccoon 

populations via anthelmintic baiting: Journal of Wildlife 

Management, v. 77, p. 1372–1379.

Smyser, T.J., Page, L.K., and Rhodes, O.E., Jr., 2010, Optimization of raccoon latrine surveys for quantifying exposure 

to Baylisascaris procyonis: Journal of Wildlife Diseases, 

v. 46, p. 929–933.

Smyser, T.J., Redding, J.V., Jr., Bevis, C.M., Page, L.K., and 

Swihart, R.K., 2015, Development of an automated dispenser for the delivery of medicinal or vaccine-laden baits 

to raccoons (Procyon lotor): Journal of Wildlife Diseases, 

v. 51, p. 513–518.

Snyder, D.E., 1983, The prevalence, cross-transmissibility to 

domestic animals and adult structure of Baylisascaris procyonis (Nematoda) from Illinois raccoons (Procyon lotor): 

Urbana, University of Illinois at Urbana-Champaign, Ph.D. 

Thesis, 233 p.

Snyder, D.E., and Fitzgerald, P.R., 1985, The relationship of 

Baylisascaris procyonis to Illinois raccoons (Procyon lotor): 

Journal of Parasitology, v. 71, p. 596–598.

Snyder, D.E., and Fitzgerald, P.R., 1987, Contaminative 

potential, egg prevalence, and intensity of Baylisascaris 

procyonis-infected raccoons (Procyon lotor) from Illinois, 

with a comparison to worm intensity: Proceedings of the 

Helminthological Society of Washington, v. 54, p. 141–145.

Sorvillo, F., Ash, L.R., Berlin, O.G.W., Yatabe, J., Degiorgio, 

C., and Morse, S.A., 2002, Baylisascaris procyonis: An 

emerging helminthic zoonosis: Emerging Infectious 

Diseases, v. 8, p. 355–359. 

Souza, M.J., Ramsay, E.C., Patton, S., and New, J.C., 2009, 

Baylisascaris procyonis in raccoons (Procyon lotor) in eastern Tennessee: Journal of Wildlife Diseases, v. 45, 

p. 1231–1234.

Sprent, J.F.A., 1951, On the migratory behavior of the larvae 

of various Ascaris species in mice: Journal of Parasitology, 

v. 37 (suppl.), p. 21.

Sprent, J.F.A., 1952a, On the migratory behavior of the larvae 

of various Ascaris species in white mice. I. Distribution of 

larvae in tissues: Journal of Infectious Diseases, v. 90, 

p. 165–176.

Sprent, J.F.A., 1952b, On an Ascaris parasite of the fisher and 

marten, Ascaris devosi sp. nov.: Proceedings of the Helminthological Society of Washington, v. 19, p. 27–37.

Sprent, J.F.A., 1953a, On the migratory behavior of the larvae 

of various Ascaris species in white mice. II. Longevity of 

encapsulated larvae and their resistance to freezing and 

putrefaction: Journal of Infectious Diseases, v. 92, 

p. 114–117.

Sprent, J.F.A., 1953b, On the life history of Ascaris devosi and 

its development in the white mouse and the domestic ferret: 

Parasitology, v. 42, p. 244–258.

Sprent, J.F.A., 1955, On the invasion of the central nervous 

system by nematodes. II. Invasion of the nervous system in 

ascariasis: Parasitology, v. 45, p. 41–55.

Sprent, J.F.A., 1968, Notes on Ascaris and Toxascaris, with a 

definition of Baylisascaris gen.nov.: Parasitology, v. 58, 

p. 185–198.

Sprent, J.F.A., 1970, Baylisascaris tasmaniensis sp.nov. in 

marsupial carnivores: heirloom or souvenir?: Parasitology, 

v. 61, p. 75–86.

Sprent, J.F.A., Lamina, J., and McKeown, A., 1973, 

Observations on migratory behaviour and development of 

Baylisascaris tasmaniensis: Parasitology, v. 67, p. 67–83.

Stains, H.J., 1956, The raccoon in Kansas: Lawrence, University of Kansas, Miscellaneous Publications of the Museum 

of Natural History, v. 10, p. 1–76.

Stefański, W., and Žarnowski, E., 1951, Ascaris procyonis n. 

sp. z jelita szopa (Procyon lotor L.) Ascaris procyonis n. sp. 

provenant de l’intestin de Procyon lotor L: Annales Musei 

Zoologici Polonici, v. 14, p. 199–202.

Stone, W.B., 1983, Intestinal obstruction in raccoons caused 

by the ascarid Baylisascaris procyonis: New York Fish and 

Game Journal, v. 30, p. 117–118.

Stringfield, C.E., and Sedgwick, C.J., 1997, Baylisascaris: A 

zoo-wide experience: Proceedings of the American Association of Zoo Veterinarians, p. 73–77.

Stubbe, M., 1990, Der Status des Waschbären Procyon lotor

(L.) in der DDR (1975–1984): Beiträge zur Jagd- und 

Wildforschung, v. 17, p. 180–192. [In German.]

Stuewer, F.W., 1943, Raccoons: Their habits and management 

in Michigan: Ecological Monographs, v. 13, p. 204–257.

Suedmeyer, W.K., Bermudez, A., and Kazacos, K.R., 1996, 

Cerebellar nematodiasis in an emu (Dromaius novaehollandiae): Journal of Zoo and Wildlife Medicine, v. 27, 

p. 544–549.

Swerczek, T.W., and Helmboldt, C.F., 1970, Cerebrospinal 

nematodiasis in groundhogs (Marmota monax): Journal of 

the American Veterinary Medical Association, v. 157, 

p. 671–674.
100 Baylisascaris Larva Migrans

Taira, K., Une, Y., Šnábel, V., and Sugiyama, H., 2013, 

Baylisascaris sp. infection in a pet kinkajou Potos flavus: 

Helminthologia, v. 50, p. 238–243.

Tarantola, R.M., Elkins, K.A., Kay, C.N., and Folk, J.C., 2011, 

Photoreceptor recovery following laser photocoagulation 

and albendazole in diffuse unilateral subacute neuroretinitis: 

Archives of Ophthalmology, v. 129, p. 669–671.

Taylor, M.R.H., Keane, C.T., O’Connor, P., Girdwood, 

R.W.A., and Smith, H., 1987, Clinical features of covert 

toxocariasis: Scandinavian Journal of Infectious Diseases, 

v. 19, p. 693–696. 

Tecec, T.G., 1987, Occurrence of Baylisascaris procyonis in 

raccoon populations on military installations in the Washington, D.C. area: Military Medicine, v. 152, p. 83–84.

Tenora, F., and Stanek, M., 1990, Scanning electron microscopy of Toxascaris procyonis Stefanski et Zarnowski, 1951 

(Nematoda): Helminthologia, v. 27, p. 73–77.

Tenora, F., Hönigová, M., and Stanek, M., 1991, Interesting 

findings of two species of Ascaridata (Nematoda)—parasites of carnivora in Czech and Slovak Federative Republic: 

Helminthologia, v. 28, p. 131–135.

Thomas, J.S., 1988, Encephalomyelitis in a dog caused by 

Baylisascaris infection: Veterinary Pathology 25: 94–95.

Thompson, A.B., Glover, G.J., Postey, R.C., Sexsmith, J.L., 

Hutchison, T.W.S., and Kazacos, K.R., 2008, Baylisascaris 

procyonis encephalitis in Patagonian conures (Cyanoliseus 

patagonus), crested screamers (Chauna torquata), and a 

western Canadian porcupine (Erethizon dorsatum epixanthus) in a Manitoba zoo: Canadian Veterinary Journal, v. 49, 

p. 885–888.

Tiner, J.D., 1949, Preliminary observations on the life history of Ascaris columnaris: Journal of Parasitology, v. 35 

(suppl.), p. 13.

Tiner, J.D., 1951, Observations on larval carnivore ascarids in 

rodents: Journal of Parasitology, v. 37 (suppl.), p. 21–22.

Tiner, J.D., 1952a, A study of the epidemiology of ascarid 

nematodes in carnivores and rodents: Urbana, University of 

Illinois, Ph.D. Thesis, 130 p.

Tiner, J.D., 1952b, Speciation in the genus Ascaris: Additional 

experimental and morphological data: Journal of Parasitology, v. 38 (suppl.), p. 27.

Tiner, J.D., 1953a, Fatalities in rodents caused by larval 

Ascaris in the central nervous system: Journal of 

Mammalogy, v. 34, p. 153–167.

Tiner, J.D., 1953b, The migration, distribution in the brain, 

and growth of ascarid larvae in rodents: Journal of 

Infectious Diseases, v. 92, p. 105–113.

Tiner, J.D., 1954, The fraction of Peromyscus leucopus fatalities caused by raccoon ascarid larvae: Journal of Mammalogy, v. 35, p. 589–592.

Tokiwa, T., Nakamura, S., Taira, K., and Une, Y., 2014, Baylisascaris potosis n. sp., a new ascarid nematode isolated 

from captive kinkajou, Potos flavus, from the Cooperative 

Republic of Guyana: Parasitology International, v. 63, 

p. 591–596.

Tokiwa, T., Tsugo, K., Nakamura, S., Taira, K., and Une, Y., 

2015, Larva migrans in squirrel monkeys experimentally 

infected with Baylisascaris potosis: Parasitology International, v. 64, p. 284–287.

Tseng, F.S., 1997, Baylisascaris in squirrels: Proceedings of 

the North American Veterinary Conference, p. 817–818.

U.S. Department of Agriculture-Animal and Plant Health 

Inspection Service, 2015, Import live animals: accessed 

September 10, 2015, at https://www.aphis.usda.gov/.

Van Andel, R.A., Franklin, C.L., Besch-Williford, C., Riley, 

L.K., Hook, R.R., Jr., and Kazacos, K.R., 1995, Cerebrospinal larva migrans due to Baylisascaris procyonis in a guinea 

pig colony: Laboratory Animal Science, v. 45, p. 27–30.

Vieira, F.M., Luque, J.L., and Muniz-Pereira, L.C., 2008, 

Checklist of helminth parasites in wild carnivore mammals 

from Brazil: Zootaxa, v. 1721, p. 1–23.

Voge, M., 1956, A list of nematode parasites from California 

mammals: American Midland Naturalist, v. 56, p. 423–429.

Waller, E.F., 1940, Infectious gastroenteritis in raccoons 

(Procyon lotor): Journal of the American Veterinary 

Medical Association, v. 96, p. 266–268.

Wang, T., He, G., Yang, G., Fei, Y., Zhang, Z., Wang, C., Yang, 

Z., Lan, J., Luo, L., and Liu, L., 2008, Cloning, expression 

and evaluation of the efficacy of a recombinant Baylisascaris schroederi Bs-Ag3 antigen in mice: Vaccine, v. 26, 

p. 6919–6924.

Wilder, H.C., 1950, Nematode endophthalmitis: Transactions 

of the American Academy of Ophthalmology, v. 55, 

p. 99–109.

Williams, C.K., McKown, R.D., Veatch, J.K., and Applegate, 

R.D., 1997, Baylisascaris sp. found in a wild northern bobwhite (Colinus virginianus): Journal of Wildlife Diseases, 

v. 33, p. 158–160.

Williams, G.A., Aaberg, T.M., and Dudley, S.S., 1988, 

Perimacular photocoagulation of presumed Baylisascaris 

procyonis in diffuse unilateral subacute neuroretinitis, in

Gitter, K.A., Schatz, H., Yannuzzi, L.A., and McDonald, 

H.R., eds., Laser photocoagulation of retinal disease: San 

Francisco, California, Pacific Medical Press, p. 275–280.
References Cited 101

Wilson, D.E., and Reeder, D.M., eds., 1993, Mammal species 

of the world: A taxonomic and geographic reference (2nd 

ed.): Washington, D.C., Smithsonian Institution Press, 

1206 p.

Windsor, R.C., Sturges, B.K., Vernau, K.M., and Vernau, W., 

2009, Cerebrospinal fluid eosinophilia in dogs: Journal of 

Veterinary Internal Medicine, v. 23, p. 275–281.

Winter, M., 2006, Procyon lotor: DAISIE, Delivering Alien 

Invasive Species Inventories for Europe, 4 p., Accessed 

May 25, 2014 at http://www.europe-aliens.org.

Winter, M., Stubbe, M., and Heidecke, D., 2005, Zur Ökologie des Waschbären (Procyon lotor L., 1758) in SachsenAnhalt: Beiträge zur Jagd- und Wildforschung, v. 30, p. 

303–322. [In German.]

Winterfield, R.W., and Thacker, H.L., 1978, Verminous 

encephalitis in the emu: Avian Diseases, v. 22, p. 336–339.

Wirtz, W.L., 1981, Host range for cerebrospinal nematodiasis 

and visceral larva migrans due to Baylisascaris procyonis: 

Proceedings of the Annual Midwestern Conference of Parasitologists, v. 33, p. 7.

Wirtz, W.L., 1982, Cerebrospinal nematodiasis due to the 

raccoon ascarid, Baylisascaris procyonis: West Lafayette, 

Indiana, Purdue University, M.S. Thesis, 90 p.

Wise, M.E., Sorvillo, F.J., Shafir, S.C., Ash, L.R., Berlin, O.G., 

2005, Severe and fatal central nervous system disease in 

humans caused by Baylisascaris procyonis, the common 

roundworm of raccoons: a review of current literature: 

Microbes and Infection, v. 7, p. 317–323. 

Wolf, K.N., Lock, B., Carpenter, J.W., and Garner, M.M., 

2007, Baylisasacaris procyonis infection in a moluccan 

cockatoo (Cacatua moluccensis): Journal of Avian Medicine 

and Surgery, v. 21, p. 220–225.

Won, K.Y., Kruszon-Moran, D., Schantz, P.M., and Jones, J.L., 

2008, National seroprevalence and risk factors for zoonotic 

Toxocara spp. infection: American Journal of Tropical 

Medicine and Hygiene, v. 79, p. 552–557.

 Wu, J., He, G., and Hu, H., 1987, Study on a new species, 

Toxascaris ailuri sp. n. (Nematoda:Ascaridae) from red 

panda: Sichuan Journal of Zoology, v. 2, p. 1–3.

Wyand-Ouellette, W.K., Kazacos, K.R., and DeNicola, D.B., 

1983, Assessment of pulmonary migration of ascarids 

(Baylisascaris, Ascaris) by bronchopulmonary lavage: 

Proceedings of the American Association of Veterinary 

Parasitologists, v. 28, p. 24.

Xie, Y., Zhang, Z., Niu, L., Wang, Q., Wang, C., Lan, J., Deng, 

J., Fu, Y., Nie, H., Yan, N., Yang, D., Hao, G., Gu, X., 

Wang, S., Peng, X., and Yang, G., 2011, The mitochondrial genome of Baylisascaris procyonis: PLoS One, v. 6, 

p. e27066 (10 pages).

Xie, Y., Zhou, X., Li, M., Liu, T., Gu, X., Wang, T., Lai, W., 

Peng, X., and Yang, G., 2014, Zoonotic Baylisascaris procyonis roundworms in raccoons, China [letter]: Emerging 

Infectious Diseases, v. 20, p. 2170–2172.

Yabsley, M., 2010, Baylisascaris in Florida raccoons: Athens, University of Georgia School of Veterinary Medicine, 

SCWDS Briefs, v. 26, p. 2–3.

Yeager, L.E., and Rennels, R.G., 1943, Fur yield and autumn 

foods of the raccoon in Illinois river bottom lands: Journal 

of Wildlife Management, v. 7, p. 45–60.

Yeitz, J.L., Gillin, C.M., Bildfell, R.J., and DeBess, E.E., 

2009, Prevalence of Baylisascaris procyonis in raccoons 

(Procyon lotor) in Portland, Oregon, USA: Journal of Wildlife Diseases, v. 45, p. 14–18.

Yoshikawa, M., Nishiofuku, M., Moriya, K., Ouji, Y., 

Ishizaka, S., Kasahara, K., Mikasa, K., Harai, T., Mizuno, 

Y., Ogawa, S., Nakamura, T., Maruyama, H., and Akao, 

N., 2008, A familial case of visceral toxocariasis due to 

consumption of raw bovine liver: Parasitology International, 

v. 57, p. 525–529.

Zhang, J.S., Daszak, P., Huang, H.L., Yang, G.Y., Kilpatrick, 

A.M., and Zhang, S., 2007, Parasite threat to panda conservation: EcoHealth , v. 5, p. 6–9.

Zimmerman, D., Dangoudoubiyam, S., and Kazacos, K.R., 

2010, Human serological testing for Baylisascaris procyonis

in nonhuman primates: Proceedings of the Joint Conference of the American Association of Zoo Veterinarians 

and American Association of Wildlife Veterinarians, South 

Padre Island, Texas, October 2010, p. 124.
102 Baylisascaris Larva Migrans

Glossary

Terms are defined relative to meanings within this publication.

A

anthelmintic A drug or other agent that kills or expels worm 

parasites, especially intestinal worms. See also parasite.

antibodies Proteins formed in the body of a vertebrate that 

are used by the immune system to identify and neutralize the 

effects of foreign invading proteins, called antigens, coming 

from bacteria, viruses, parasites, and their enzymes or other 

proteins. See also antigens, parasite, vertebrate.

antigens Foreign proteins and other substances that, upon 

entering the body, stimulate the host’s immune system to 

produce protective antibodies and immune cells as a means of 

protecting the host from harmful invasion. See also antibodies, 

host.

ascarid Large roundworms in the taxonomic phylum Nematoda that live in the intestines of vertebrates; order Ascaridida, 

superfamily Ascaridoidea. Includes Ascaris, Baylisascaris, 

Toxocara, and others. See also nematode, taxonomy, vertebrate.

asymptomatic Without symptoms, or producing no symptoms. See also symptoms.

ataxia An inability to coordinate muscle activity during voluntary movement; may involve the limbs, head, or trunk.

B

Baermann funnel A funnel apparatus used to isolate parasite 

larvae from animal tissues or soil, based on the principle of 

continued activity and penetration through mesh in a warm 

solution (saline or water). See also larva, parasite.

baylisascariasis Infection or disease caused by nematodes 

in the genus Baylisascaris; usually involves larval infection 

(visceral, ocular, neural larva migrans). See also genus, larva, 

nematode, viscera, neural larva migrans (NLM), ocular larva 

migrans (OLM).

bear Any of the carnivorous or omnivorous mammals of 

the family Ursidae, having massive bodies, coarse heavy fur, 

relatively short limbs, almost rudimentary tails, and walking 

with the entire sole and heel of the foot on the ground. See 

also carnivorous, mammals, omnivorous, taxonomy.

birds Warm-blooded vertebrates with wings and feathers, 

belonging to the class Aves. The wings are poorly developed 

for some species of flightless birds. See also vertebrate.

C

cache, caching To hoard, stockpile, store up, collect. Used 

to refer to the behavior of rodents such as the Allegheny 

woodrat and other packrats that habitually collect items and 

store them in their middens. See also midden, rodent, woodrat.

cacomistle A carnivorous procyonid mammal, similar in 

appearance to the ringtail and found from southern Mexico to 

western Panama. See also carnivorous, mammals, procyonid, 

ringtail.

canids Carnivorous mammals within the family Canidae (for 

example, wolves, coyotes, foxes, and other doglike animals). 

See also carnivorous, taxonomy.

carnivore, carnivorous A flesh-eating mammal of the order 

Carnivora, comprising the wolves, bears, raccoons, cats, weasels, etc. Their teeth are adapted for feeding on flesh. See also

bear, canids, mammals, raccoon.

central nervous system (CNS) The part of the nervous system comprising the brain and spinal cord.

cerebrospinal Relating to the brain and the spinal cord.

cerebrospinal fluid (CSF) The fluid in the ventricles of the 

brain and surrounding the spinal cord. See also cerebrospinal.

cerebrospinal nematodiasis Invasion of the brain and (or) 

spinal cord by nematode larvae. An older term referring only 

to nematodes, this has been replaced by the broader classification called neural larva migrans. See also cerebrospinal, larva, 

nematode, neural larva migrans. 

clinical Readily observable.

clinical sign Objective evidence of disease or injury that can 

be perceived by an examiner. See also clinical.

coati A tropical American carnivore of the genus Nasua, 

related to the raccoon, having an elongated body, long, ringed 

tail, and a slender, flexible snout. See also carnivore, genus, 

raccoon, taxonomy.

coma, comatose A state of deep unconsciousness from which 

a person or animal cannot be aroused. Such individuals cannot 

react to events in their environment.

communicable disease An illness caused by a specific infectious agent or its toxic products, arising through transmission 

of that agent or its products from an infected person, animal, 

or inanimate reservoir directly or indirectly (via intermediate plant or animal host) to a susceptible host. See also host, 

intermediate host.

companion animals Animals maintained by humans as pets 

(such as dogs, cats, captive wildlife, horses, etc.).

computed tomography (CT) Also known as computerized 

axial tomography, an imaging process that demonstrates various bodily structures based on their ability to block an x-ray 

beam, using a computer to manipulate (“window”) the volume 

of data that is generated by the scanner.

conure Any of several small, long-tailed New World parrots, 

especially of the genus Aratinga, certain species of which are 

often kept as pets. Also referred to as large parakeets. See also

genus, taxonomy.

cortical Relating to the outer layer of the cerebrum, the part 

of the brain which coordinates the body’s sensory and motor 

information.
Glossary 103

epidemiology The study of the causes, occurrence, and control of disease in populations.

excretory-secretory (ES) antigens Metabolic antigens (proteins) excreted or secreted by living parasites such as helminth 

larvae during migration or feeding. These antigens include 

digestive enzymes, nitrogenous and other waste products, 

proteins shed from the surface, and other substances. Immune 

responses of the host are often directed against ES antigens 

produced by migrating larvae. See also antigens, helminth, 

host, larva, parasite.

exotic pet A pet of foreign origin or character; not a native 

species, but introduced from abroad.

F

fecal examination Examination of samples of feces for 

infectious and parasitic agents. Usually refers to examination 

for parasite eggs, larvae, cysts, or other stages. See also feces, 

larvae, parasite.

fecal flotation A method of examining samples of feces in 

which parasite stages (eggs, larvae, cysts) are isolated from 

feces by using flotation on high density solutions, usually various salts or sugar. See also feces, larvae, parasite.

feces Waste matter discharged from the intestines through 

the anus; excrement.

G

general practitioner A medical professional, such as a 

physician or veterinarian, whose practice is not limited to any 

specific branch of medicine or class of diseases.

genus, genera A taxonomic group of similar species; similar 

genera are grouped into families. See also species, taxonomy.

geophagia Eating earthy matter, especially soil or sand.

granuloma An inflammatory nodule, tumor or growth 

composed of connective tissue and other cells, often forming 

around helminth (worm) larvae or other organisms in body 

tissues. See also helminth, larva. 

H

helminth A worm or wormlike organism. In a medical context, refers to worm parasites, including nematodes 

(roundworms), cestodes (tapeworms), trematodes (flukes), and 

acanthocephalans (thorny-headed worms). See also parasite.

hemorrhage A profuse discharge of blood, as from a ruptured blood vessel; bleeding.

histology, histological The study of the microscopic structure of tissues.

histopathology The science dealing with the histological 

structure of abnormal or diseased tissue; pathological histology. See also histology, pathological.

covert Something that is concealed, hidden, secret, or disguised. A covert infection is one that is present but with nonspecific clinical signs that cannot be specifically diagnosed. 

Covert infections may be detected by isolating the agent or 

its antigens or by detecting specific antibodies in the serum. 

See also antibodies, antigens, clinical sign, serum.

D

definitive host A host in which a parasite reaches the adult 

stage or undergoes sexual reproduction. See also host, parasite.

dementia A pattern of mental decline caused by different 

diseases or conditions, such as brain injury, infection, degenerative diseases, adverse drug reactions, etc.

developmental delay The developmental condition wherein 

a child does not demonstrate abilities and skills commonly 

found in other children in the same age group. Developmental delays may involve physical abilities (gross or fine motor 

skills), language, or personal or social skills.

diffuse unilateral subacute neuroretinitis (DUSN) A type of 

ocular larva migrans resulting in inflammatory and degenerative changes to the retina, retinal vessels, and optic nerve head, 

caused by larvae of Baylisascaris, Toxocara, Ancylostoma, 

Alaria, and probably other parasites. See also inflammation, 

ocular larva migrans, parasite. 

DNA Deoxyribonucleic acid, a nucleic acid found mainly in 

the chromosomes of an organism that contains the hereditary 

information of that organism.

E

emaciated Abnormally thin, caused by lack of nutrition or 

by disease.

encephalitis Inflammation of the substance of the brain.

enzootic Occurring commonly within an animal population 

or a geographical area.

enzyme-linked immunosorbent assay (ELISA) A laboratory 

test used to detect the presence of either antigen or antibody 

in a sample using an enzyme reagent that generates a color 

reaction.

eosinophil A white blood cell involved in combating parasitic and certain other infections and also involved in allergic 

reactions. See also eosinophilia, helminth, parasite.

eosinophilia An abnormally increased number of eosinophils, a type of white blood cell, in the blood. See also eosinophil.

eosinophilic meningitis or meningoencephalitis A type of 

inflammation of the brain and its coverings in which white 

blood cells called eosinophils are the predominant cell type; 

eosinophils usually increase in response to specific antigens 

produced by helminth larvae, fungi, or other agents. See also

antigen, eosinophil, helminth, inflammation, parasite.
104 Baylisascaris Larva Migrans

host A living plant or animal that harbors and (or) nourishes microbes, viruses, or parasites. See also definitive host, 

paratenic host, parasite.

I

indirect immunofluorescence A diagnostic method using a 

fluorescence microscope to detect specific antigens (proteins, 

etc.) in tissues based on their binding of antibodies carrying 

fluorescent dyes. See also antibodies, antigens.

infectious disease A disease resulting from infection with a 

pathogenic microbial agent. See also pathogenic.

inflammation Redness, swelling, pain, tenderness, heat, and 

disturbed function of an area of the body, especially as a reaction of tissues to injurious agents.

intermediate host A host in which a parasite undergoes 

essential development of intermediate or larval stages but not 

to sexual maturity. See also host, parasite.

invertebrates Animals lacking a spinal column, for example, 

insects and helminth (worm) parasites; vertebrates, in contrast, 

possess a spinal column. See also helminth.

isolate, isolation To separate something of interest from 

surrounding material in order to facilitate further study and 

identification.

J

K

kinkajou An arboreal carnivore with a prehensile tail, Potos 

flavus, of Central and South America, related to the raccoon 

and coati. See also carnivore, coati, raccoon.

L

larva, larvae, larval An immature stage of an organism that 

is fundamentally different from the adult.

larva migrans (LM) The prolonged migration and persistence 

of helminth larvae in the organs and tissues of humans and 

animals. See also helminth, larva, parasite.

larvicidal Pertaining to an agent, such as an anthelmintic 

drug, with the ability to kill larvae. See also anthelmintic.

lesion An abnormal change in the appearance of tissue or 

loss of normal function due to pathogens, injury, or other 

causes. See also pathogen.

M

magnetic resonance imaging (MRI) An imaging method 

that detects various bodily structures in great detail by using 

powerful magnetic fields to align the magnetization of atomic 

nuclei in cells in the tissues.

mammals Warm-blooded vertebrate animals that possess 

hair during some part of their life and suckle their young. 

See also vertebrate.

midden A dunghill, debris, or garbage pile, or a pile of 

refuse. Rodents such as ground squirrels and woodrats may 

use burrows or other areas for food storage, and in the case 

of woodrats, also known as packrats, various items, including food material, animal bones, fecal material, and disparate 

shiny or interesting objects, may be cached in these large 

debris piles. See also cache, rodent, woodrat.

N

necropsy Examination and dissection of an animal after 

death, usually to determine a cause of death; postmortem 

examination.

nematode Unsegmented, cylindrical parasitic worm of the 

class Nematoda; roundworm.

neural larva migrans (NLM) Invasion of the brain and (or) 

spinal cord by helminth larvae, such as nematodes in the genera Baylisascaris and Toxocara. The raccoon ascarid, B. procyonis, is a well-known cause of NLM in humans and animals. 

See also ascarid, genus, helminth, nematode, parasite, raccoon.

neurologic disease A disease affecting the nervous system 

(brain, spinal cord, etc.).

neurologist A medical professional, such as a physician or a 

veterinarian, who specializes in diseases and conditions of the 

nervous system.

neurotropic Having a predilection for the nervous system or 

nervous tissue.

O

ocular larva migrans (OLM) Invasion of the eye by helminth 

larvae, such as nematodes in the genera Baylisascaris and Toxocara. See also genus, helminth, larvae, nematode, parasite.

omnivorous Having a diet of both plants and animals.

ophthalmologist A medical professional, such as a physician 

or a veterinarian, who specializes in diseases of the eye.

P

parasite, parasitic An organism that lives at least part of its 

life cycle on or within another organism (the host) upon which 

it is physiologically dependent and to which it causes harm 

in some way. In a parasitologic or medical context, it usually refers to animal agents, that is, protozoa, helminths, and 

arthropods. See also helminth, host.

paratenic host A host in which a parasite survives without 

undergoing required or essential development. A paratenic 

host accumulates and maintains intermediate or larval stages 

of a parasite which may undergo growth, but is not essential or 

required in the parasite’s life cycle. Also known as a transport 

host. See also host, parasite, paratenesis.

patent Refers to a mature parasitic infection that results in 

the appearance of parasite eggs or larvae in or from the host. 

See also host, larva, parasitic, prepatent period.

pathogen, pathogenic Any organism capable of producing 

disease; typically refers to microorganisms (bacteria, viruses, 
Glossary 105

protozoa), but broadly includes all disease-producing agents, 

including worm parasites. See also parasite.

pathogenicity The ability of an organism to cause disease.

pathology The study of the structural and functional effects 

of disease.

pathological Pertaining to pathology. See also clinical sign, 

histopathology, pathology, symptoms.

pediatrician A physician who specializes in the diseases of 

children.

peridomestic Around or in the vicinity of human habitation.

pheasant Long-tailed and often brightly colored Old World 

gallinaceous birds of the family Phasianidae. See also birds.

pharmacokinetics The absorption, movement, and other 

characteristics of distribution and elimination of drugs from 

the body.

phylogenetics The study of evolutionary relatedness among 

groups of organisms (such as species and populations) using 

molecular sequencing and morphological data, useful in biological systematics and taxonomy.

pica An eating disorder characterized by compulsive craving 

and eating of nonnutritive substances that may include clay, 

dirt, sand, chalk, paper, etc.

polymerase chain reaction (PCR) A molecular biologic 

technique that uses an enzyme (deoxyribonucleic acid [DNA] 

polymerase) and short specific DNA fragments (primers) to 

enable selective and repeated amplification of a single or a few 

copies of a piece of DNA across many orders of magnitude, 

generating thousands to millions of copies of a particular DNA 

sequence, which can then be detected using gel electrophoresis. The method has become very important and widespread 

for the detection and diagnosis of pathogens in bodily tissues, 

fluids and other samples. See also DNA.

population A group of organisms inhabiting a specific area or 

possessing a specific trait.

poultry A general term for farmed domestic birds, including 

chickens, turkeys, ducks, and geese. See also birds.

predator An animal that captures, kills, and eats other animals for food.

prepatent period The period of time from initial infection with a parasite until one can demonstrate that the host is 

infected by finding stages of the parasite (eggs, larvae, etc.) 

in or from the host; also known as the biological incubation 

period. The patent period is the period of time from first demonstration of a parasitic infection (by finding eggs, larvae, etc.) 

until one can no longer demonstrate that the host is infected. 

See also host, larvae, parasite, patent.

prevalence The total number of cases of a disease in a 

population at a given time. Also, the proportion of individuals found to be positive (infected) for a pathogen or parasite 

of those individuals tested or examined. See also pathogen, 

parasite, population.

primate Any of various omnivorous mammals of the order 

Primates, comprising humans, the apes, Old and New World 

monkeys, lemurs, and tarsiers, that are especially distinguished 

by the use of hands, varied locomotion, and complex social 

behavior. See also omnivore, taxonomy.

procyonid A mammal within the taxonomic family Procyonidae. They are relatively small, omnivorous New World 

mammals including raccoons, coatis, kinkajous, olingos, and 

ringtails. Most procyonids have distinct masklike facial markings and banded tails. See also coati, mammals, omnivorous, 

raccoon, ringtail, taxonomy.

pseudoparasite From a diagnostic standpoint, nonparasite 

material or artifacts (for example, pollen, spores, plant hairs, 

etc.) that are mistaken for parasite stages.

Q

quail Various small, short-tailed gallinaceous birds of the 

family Phasianidae. See also birds, taxonomy.

R

rabbit Small, long-eared mammals in the family Leporidae 

(rabbits and hares) of the order Lagomorpha (rabbits, hares, 

and pikas). See also mammals, taxonomy.

raccoon A nocturnal carnivore, Procyon lotor, in the family 

Procyonidae, having a masklike black stripe across the eyes, a 

sharp snout, and a bushy, ringed tail, native to North and Central America. See also carnivore, procyonid, taxonomy.

raccoon latrine An area of communal defecation by raccoons, where their feces accumulate, sometimes in substantial 

amounts. Often found on logs, large rocks, at the bases of 

trees, in barns and other outbuildings, and on rooftops.

raptor A predatory bird such as a hawk, eagle, and owl, with 

bill and claws adapted for seizing prey. See also birds.

ratite A member of a diverse group of large, flightless birds, 

including ostrich, emu, rhea, cassowary, and kiwi.

recombinant antigen A protein produced using recombinant 

DNA technology. See also antigens, recombinant DNA.

recombinant DNA DNA produced by combining genetic 

material from two or more sources by means of genetic engineering. Often used to express diagnostic or other proteins 

encoded by DNA sequences by cloning them into bacteria or 

yeasts. See also DNA.

ringtail A carnivorous procyonid mammal, Bassariscus 

astutus, of Mexico and the southwestern United States, related 

to the raccoon but smaller, with a sharper snout and longer, 

bushy ringed tail. See also carnivorous, mammals, procyonid.

rodent A diverse group of mammals characterized by incisor teeth that grow throughout life. Species within the order 

Rodentia include mice, rats, squirrels, chipmunks, voles, 
106 Baylisascaris Larva Migrans

gophers, marmots, beavers, porcupines, and many others. See 

also mammals, mice, species, taxonomy, vole.

S

scavenger An animal, such as a vulture, crow, hyena, and 

jackal, that feeds on carcasses, other carrion, and refuse.

serology Laboratory evaluations of the serum portion of 

the blood for the purpose of detecting and measuring host 

antibody levels and responses to infectious agents and other 

antigens. See also antibodies, host, serum.

seropositive The presence of an antibody specific to an 

infectious agent or other antigen in the blood as determined by 

a laboratory test. See also antibodies, antigens, serum.

serum The pale fluid that remains after blood has clotted. It 

contains a large amount and variety of proteins including antibodies and other substances. See also antibodies, seropositive.

shrew A small molelike omnivorous mammal in the order 

Soricomorpha, family Soricidae, possessing sharp, spikelike 

teeth, poor eyesight, and excellent senses of hearing and smell. 

They are very active, with voracious appetites and unusually 

high metabolic rates. See also mammals, omnivorous, taxonomy.

skunk Several species of carnivores within the family 

Mephitidae well-known for their pungent anal glands; specifically, the striped, hooded, spotted, and hog-nosed skunk. See 

also carnivore, species.

somatic (tissues, migration) Pertaining to the body wall 

proper (muscles, connective tissue, etc.) as opposed to the 

viscera. See also viscera.

species A population of organisms whose members are 

able to breed among themselves and produce fertile offspring. 

More precise determinations of species are based on similarity 

of deoxyribonucleic acid (DNA). See also DNA, population, 

taxonomy.

specificity The restriction of a parasite to one or more species of hosts, that is, the degree to which a parasite is able to 

develop in more than one host species. See also host, parasite, 

species.

squirrel Small to medium-sized arboreal rodents in the family Sciuridae, having long, bushy tails and strong hind legs.

stage Any particular form in the life cycle of a parasite 

that can be distinguished from all of its other forms. See also

parasite.

stupor, stuporous A state of impaired consciousness in 

which the patient shows a marked diminution in reactivity to 

environmental stimuli and can be aroused only by continual 

stimulation.

subclinical A condition without overt clinical manifestations, yet sometimes able to be detected using specialized 

equipment or procedures. See also clinical.

symptoms Subjective indications of a disorder or disease, 

experienced by an individual as a change from normal function, sensation, or appearance.

T

taxonomic, taxonomy The systematic principles and procedures of grouping and arranging organisms into a hierarchical 

framework (below).

Taxonomic hierarchy, 

from general to specific

Kingdom

Phylum

Class

Order

Family

Subfamily

Genus

Species

Toxocara A genus of large roundworms or ascarids (nematodes) that includes species infecting dogs, cats, foxes, and 

other carnivores. See also ascarid, carnivore, nematode, taxonomy, toxocariasais, toxocaral.

toxocaral Relating to infection or disease caused by Toxocara roundworms. See also Toxocara.

translocation Human capture of wildlife at one geographic 

area and their transportation and release at a different geographic area.

transplacental Transmitted across the placenta. Some 

infectious agents and parasites are transmitted from mother to 

offspring in utero across the placenta. See also parasite.

U

ungulate A hoofed, grazing mammal.

V

viscera, visceral The organs in the cavities of the body, 

especially those in the abdominal cavity; pertaining to those 

organs.

visceral larva migrans (VLM) Invasion of the viscera (liver, 

lungs, etc.) by helminth larvae, such as nematodes in the 

genera Baylisascaris and Toxocara. See also helminth, larvae, 

nematode, parasite, viscera, Toxocara.

voles Small rodents of the genus Microtus and related genera 

that typically have a stout body, rather blunt nose (compared 

to mice), and small ears. See also genus, mice, rodent.

W

waterfowl Birds within the family Anatidae, including species of ducks, geese, and swans. See also birds, taxonomy.
Glossary 107

Western blotting A laboratory diagnostic format used to 

detect reactivity to specific proteins in a tissue homogenate, 

extract, or complex mixture by separating the native proteins 

by gel electrophoresis, transferring them to a membrane, reacting the separated proteins with serum (that is, antibodies), and 

detecting resultant antigen-antibody reactions using a secondary antibody linked to a reporter enzyme that drives a colorimetric reaction. See also antibody, antigen, serum.

woodrat Medium to large rodents (order Rodentia) in 

the genus Neotoma, also known as packrats. Have a ratlike 

appearance with long tails, large ears, and large black eyes, 

and a characteristic behavior of gathering food material, 

bones, feces, and other objects and caching them in middens in 

or near their den. See also cache, feces, genus, midden, rodent, 

taxonomy.

X

Y

Z

zoonosis An infection or disease of animals that can be 

transmitted to humans.

zoonotic Of or relating to or constituting a zoonosis. See 

also zoonosis.

Appendixes

Appendix 1. Common and scientific names of animals cited.

Appendix 2. Cases of Baylisascaris neural larva migrans in pets.

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.
110 Baylisascaris Larva Migrans

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Mammals

Allegheny woodrat Neotoma magister

American badger Taxidea taxus

American beaver Castor canadensis

American red squirrel Tamiasciurus hudsonicus

Bennett’s wallaby Macropus rufogriseus rufogriseus

Black-and-white ruffed lemur Varecia variegata

Black-mantled tamarin Saguinus nigricollis

Black-tailed prairie dog Cynomys ludovicianus

Bornean orangutan Pongo pygmaeus

Botta’s pocket gopher Thomomys bottae

Brazilian porcupine Coendou prehensilis

Brown rat (or laboratory rat) Rattus norvegicus

Cacomistle Bassariscus sumichrasti

California ground squirrel Otospermophilus beecheyi

California pocket mouse Chaetodipus californicus

Capybara Hydrochoerus hydrochaeris

Coati Nasua narica, N. nasua

Common squirrel monkey Saimiri sciureus

Coquerel’s giant mouse lemur Mirza coquereli

Cottontop tamarin Saguinus oedipus

Coyote Canis latrans

Cozumel raccoon Procyon pygmaeus

Crab-eating macaque Macaca fascicularis

Crab-eating raccoon Procyon cancrivorus

De Brazza’s monkey Cercopithecus neglectus

Deer mouse Peromyscus maniculatus

Desert cottontail Sylvilagus audubonii

Diana monkey Cercopithecus diana

Domestic cat Felis catus

Domestic cow Bos taurus

Domestic dog Canis lupus familiaris
Appendix 1 111

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Mammals—Continued

Domestic ferret (domesticated European polecat) Mustela putorius furo

Domestic pig Sus scrofa domesticus

Domestic sheep Ovis aries 

Douglas squirrel Tamiasciurus douglasii

Dusky-footed woodrat Neotoma fuscipes

Eastern chipmunk Tamias striatus

Eastern cottontail Sylvilagus floridanus

Eastern fox squirrel Sciurus niger

Eastern gray squirrel Sciurus carolinensis

Emperor tamarin Saguinus imperator

European rabbit (domesticated) Oryctolagus cuniculus

Fisher Martes pennati

Francois’ langur Trachypithecus francoisi

Giant panda Ailuropoda melanoleuca

Golden hamster Mesocricetus auratus

Golden-headed lion tamarin Leontopithecus chrysomelas

Guinea pig (domesticated) Cavia porcellus

Hamadryas baboon Papio hamadryas

Hispid cotton rat Sigmodon hispidus

House mouse (or laboratory mouse) Mus musculus

Human Homo sapiens

Japanese macaque Macaca fuscata fuscata

Kinkajou Potos flavus

Least weasel Mustela nivalis

Lion-tailed macaque Macaca silenus

Long-nosed potoroo Potorous tridactylus

Long-tailed chinchilla Chinchilla lanigera

Long-tailed weasel Mustela frenata

Mantled guereza Colobus guereza kikuyuensis

Marten Martes americana

Meadow jumping mouse Zapus hudsonius

Meadow vole Microtus pennsylvanicus
112 Baylisascaris Larva Migrans

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Mammals—Continued

Mohol bushbaby Galago moholi

Mongolian gerbil Meriones unguiculatus

Mountain beaver Aplodontia rufa

Muskrat Ondatra zibethicus

North American porcupine Erethizon dorsatum

Northern olingo Bassaricyon gabbii

Northern short-tailed shrew Blarina brevicauda

Northern white-cheeked gibbon Nomascus leucogenys

Nutria Myocastor coypus

Olive baboon Papio anubis

Patagonian mara Dolichotis patagonum

Prairie vole Microtus ochrogaster

Prevost’s squirrel Callosciurus prevostii

Quoll Dasyurus spp.

Raccoon Procyon lotor

Red fox Vulpes vulpes

Red kangaroo Macropus rufus

Red ruffed lemur Varecia rubra

Red-handed or Midas tamarin Saguinus midas

Red-tailed squirrel Sciurus granatensis

Rhesus macaque Macaca mulatta

Ringtail Bassariscus astutus

Ring-tailed lemur Lemur catta

Rodrigues flying fox Pteropus rodricensis

Siamang Symphalangus syndactylus

Southern flying squirrel Glaucomys volans

Southern sea otter Enhydra lutris nereis

Spider monkey Ateles spp.

Striped skunk Mephitis mephitis

Sumatran orangutan Pongo abelii

Tasmanian devil Sarcophilus harrisii

Thirteen-lined ground squirrel Ictidomys tridecemlineatus
Appendix 1 113

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Mammals—Continued

Tiger quoll Dasyurus maculatus

Townsend’s mole Scapanus townsendii

Virginia opossum Didelphis virginiana

Western gray squirrel Sciurus griseus

Western harvest mouse Reithrodontomys megalotis

Western lowland gorilla Gorilla gorilla gorilla

Western pocket gopher Thomomys mazama

White-eared titi monkey Callicebus donacophilus

White-footed mouse Peromyscus leucopus

White-handed gibbon Hylobates lar

White-headed lemur Eulemur albifrons

White-headed or Geoffroy’s marmoset Callithrix geoffroyi

White-tailed deer Odocoileus virginianus

Wolf Canis lupus

Woodchuck Marmota monax

Woylie (brush-tailed bettong) Bettongia penicillata

Yellow-footed rock wallaby Petrogale xanthopus

Birds

African grey parrot Psittacus erithacus

American crow Corvus brachyrhynchos

American robin Turdus migratorius

Australian brushturkey Alectura lathami

Barn owl Tyto alba

Black-capped chickadee Parus atricapillus

Black-crowned night heron Nycticorax nycticorax

Blue jay Cyanocitta cristata

Blue-and-yellow or blue-and-gold macaw Ara ararauna

Blue-crowned parakeet or conure Aratinga acuticaudata

Blue-fronted Amazon Amazona aestiva

Blue-naped mousebird Urocolius macrourus

Brown thrasher Toxostoma rufum

Budgerigar Melopsittacus undulatus
114 Baylisascaris Larva Migrans

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Birds—Continued

Burrowing parrot Cyanoliseus patagonus

Bushtit Psaltriparus minimus

California quail Callipepla californica

California thrasher Toxostoma redivivum

Carolina wren Thryothorus ludovicianus

Chukar Alectoris chukar

Cockatiel Nymphicus hollandicus

Common canary Serinus canaria

Common pheasant Phasianus colchicus

Cuban Amazon Amazona leucocephala

Dark-eyed junco Junco hyemalis

Diamond dove Geopelia cuneata

Domestic canary Serinus canaria domestica

Domestic chicken Gallus gallus domesticus

Domestic duck Anas platyrhynchos domesticus

Downy woodpecker Picoides pubescens

Eastern rosella Platycercus eximius

Eastern screech owl Otus asio

Emu Dromaius novaehollandiae

European starling Sturnus vulgaris

Galah Eolophus roseicapilla

Greater rhea Rhea americana

Greater roadrunner Geococcyx californianus

Green-naped lorikeet Trichoglossus haematodus haematodus

Helmeted guineafowl Numida meleagris

Hermit thrush Catharus guttatus

House finch Carpodacus mexicanus

House sparrow Passer domesticus

House wren Troglodytes aedon

Hybrid blue-and-yellow X Scarlet macaw 

(“Catalina” macaw)

Ara ararauna X A. macao

Inca tern Larosterna inca

Indian peafowl Pavo cristatus
Appendix 1 115

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Birds—Continued

Loggerhead shrike Lanius ludovicianus

Lovebirds Agapornis spp.

Mallard X Anas platyrhynchos X

Marigold (Edward’s) lorikeet Trichoglossus capistratus

Military macaw Ara militaris

Mourning dove Zenaida macroura

Northern bobwhite Colinus virginianus

Northern cardinal Cardinalis cardinalis

Northern flicker Colaptes auratus

Northern mockingbird Mimus polyglottos

Northern red-billed hornbill Tockus erythrorhynchus

Orange-fronted parakeet or conure Aratinga canicularis

Ornate lorikeet Trichoglossus ornatus

Ostrich Struthio camelus

Ovenbird Seiurus aurocapillus

Rainbow lorikeet Trichoglossus haematodus 

Red-and-green macaw Ara chloropterus

Red-crowned (green-cheeked) Amazon Amazona viridigenalis

Red-tailed black cockatoo Calyptorhynchus banksii

Rock partridge Alectoris graeca

Rock pigeon Columba livia

Rosy-faced lovebird Agapornis roseicollis

Ruffed grouse Bonasa umbellus

Salmon-crested or moluccan cockatoo Cacatua moluccensis

Sanderling Calidris alba

Scarlet macaw Ara macao

Southern screamer Chauna torquata

Speckled mousebird Colius striatus

Spotted towhee Pipilo maculatus

Sun parakeet or conure Aratinga solstitialis

Swainson’s lorikeet Trichoglossus haematodus moluccanus

Swainson’s thrush Catharus ustulatus
116 Baylisascaris Larva Migrans

Appendix 1. Common and scientific names of animals cited.—Continued

Common name Scientific name

Birds—Continued

Thick-billed parrot Rhynchopsitta pachyrhyncha

Tufted titmouse Baeolophus bicolor

Western scrub jay Aphelocoma californica

White (umbrella) cockatoo Cacatua alba

White-breasted nuthatch Sitta carolinensis

Wild turkey Meleagris gallopavo

Wood thrush Hylocichla mustelina

Yellow-backed lorikeet Lorius garrulus flavopalliatus

Yellow-headed Amazon or yellow-headed parrot Amazona oratrix

Yellow-naped Amazon Amazona auropalliata

Yellow-rumped warbler Dendroica coronata

Roundworms (nematodes)

Dog roundworm Toxocara canis

Raccoon roundworm Baylisascaris procyonis

Hookworms (nematodes)

Dog and cat hookworms Genus Ancylostoma

Flatworms or flukes (trematodes)

Intestinal flukes of carnivores Genus Alaria

Insects

Bed bugs Genus Cimex
Appendix 2 117

Appendix 2. Cases of Baylisascaris neural larva migrans in pets.

[NR, not reported or details unavailable; >,greater than]

Animal(s) Location

Deaths or 

animals 

affected per 

total number 

of animals

Source of infection Reference

Dog Michigan 1 of 1 The owner also had a raccoon that shed roundworms 

in its feces. A puppy was exposed to the raccoon 

and its environment.

Thomas (1988).

Dog Indiana 1 of 3 A puppy was housed with two littermates in a building that also housed a raccoon in a wire cage. 

Puppies had access to an area around the raccoon 

cage that was littered with feces and food.

Rudmann and others (1996).

Dog California 1 of 1 A dog had access to a small property known to 

contain raccoon latrines.

H.R. Galano and others, 

unpub. data (2005).

Rabbits Indiana 2 of 2 The owner housed a wild young raccoon for 8 

months in the same cage until 3 months prior to 

using it for the rabbits. Raccoon feces were still 

present in the cage when it was used for the 

rabbits. 

D.W. Knapp and K.R. Kazacos, 

unpub. data (1985).

Rabbit Indiana 1 A pet rabbit was placed in a cage that 2 years previously had housed two juvenile raccoons for rehabilitation. At no time did the raccoons receive 

any treatments for parasites.

J.A. Engelhardt, 

unpub. data (1987).

Rabbits Washington 4 of 4 Rabbits were kept by two owners and had access to 

an outdoor enclosure and a yard. Raccoons were 

common in the area and often were seen by the 

owners.

Deeb and DiGiacomo (1994).

Rabbits Illinois 2 of 2 House rabbits were let out in the backyard in 

Chicago where raccoons were common.

K.R. Kazacos, unpub. data 

(1996).

Golden 

hamster

Washington 1 of 1 NR M.M. Garner, written commun. 

(2010).

North 

American 

porcupines

Indiana 2 of 2 Two pet porcupines were housed in a cage 

previously occupied by raccoons. The porcupines 

also had access to the ground area around the 

cage. Soil samples around the cage contained 

infective B. procyonis eggs.

Fitzgerald and others (1991).

Cockatiels Iowa 3 of 3 The birds were briefly housed in a cage previously 

used for two young raccoons.

Myers and others (1983).

Canaries California 12 of 22 Twenty-two canaries were housed in an outdoor 

aviary. Raccoons were commonly seen near the 

aviary.

Loretti and others (2008).

Cockatoo Kansas 1 of 1 A cockatoo owned by a wildlife rehabilitator was 

intermittently exposed to a cage previously used 

to house juvenile raccoons.

Wolf and others (2007).

Spider 

monkey

Massachusetts 1 of 1 A spider monkey was housed in an area where 

raccoons had been kept previously.

Garlick and others (1996).

Parrots and 

parakeets

Indiana 10 of 10 The birds were fed a seed mixture that was stored in 

a person’s garage and became contaminated 

by wild raccoons that had access to the area.

A.M. Lennox and others, 

unpub. data (1996); 

Lennox and others (2015).

Scarlet 

macaws

Iowa 2 of 2 NR M.A. Nieves and others, 

unpub. data (1989).

Pigeons Nebraska >15 of 200 The birds were fed a seed mixture containing sorghum that was contaminated with raccoon feces.E.N. Pendleton and V.W. Rinne, 

unpub. data (1992).
118 Baylisascaris Larva Migrans

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.—Continued

[CNS, central nervous system; NR, not reported; >, greater than]

Animal(s) Location Facility type

Number of 

deaths or 

animals 

affected

Source of infection Reference

Rabbits Michigan Commercial 

rabbitry

80 Although the source was not determined, hay and 

straw bedding used for the rabbits were stored in 

a barn loft that was accessible by wild birds and 

mammals; raccoons or skunks were suspected.

Dade and others 

(1975).

Rabbits Indiana Private 

rabbitry

20–25 Twenty to 25 rabbits were kept in outdoor hutches, 

where the bedding consisted of contaminated 

straw stored in a barn used by raccoons as a den 

and latrine site.

Kazacos and others 

(1983).

Rabbits Indiana Private 

rabbitry

20 Raccoons were common on the farm where the 

rabbits were kept and had access to the rabbit 

feed. A raccoon had been kept in one of the rabbit 

cages approximately 2 months prior.

D.D. Harrington, 

unpub. data 

(1982).

Chickens Indiana Farm 622 A flock of 11,000 chickens was housed on straw litter in a confinement house. The straw litter for the 

birds came from a barn contaminated by raccoons. 

B. procyonis eggs were found in fecal samples 

from the straw.

Richardson and 

others (1980).

Bobwhites Indiana Farm 85 Eighty-five bobwhite quail had access to a dirt run 

that had previously housed three young raccoons. 

Huge numbers of infective B. procyonis eggs were 

found in soil samples from the pen, and feces of 

the raccoons were positive.

Reed and others 

(1981); Kazacos and others 

(1982).

Bobwhites Illinois Farm 640 A flock of 712 bobwhite quail were exposed to pens 

and straw contaminated by wild raccoons.

K.R. Kazacos and 

others, unpub. 

data (2002).

Emus California Farm 5 Raccoons and latrines were present near the area 

where 18 emus were fed.

Loretti and others 

(2008).

Long-tailed 

chinchillas

Ontario Ranch 100 One hundred chinchillas from three ranches developed neurological disease. Hay used on the 

ranches came from a barn where 

raccoons had denned, and raccoon feces were 

noted on the hay.

Sanford (1989).

Long-tailed 

chinchillas

Minnesota Farm 17 At a farm where 1,400 chinchillas were housed, 

low-level exposure to B. procyonis eggs through 

contaminated hay was suspected in the deaths.

A.Wuenschmann 

and others, unpub. data (2007).

Emus Ontario Farm 9 chicks

2 adults

Breeder birds and 35 emu chicks were housed in 

indoor or outdoor pens with straw bedding from a 

barn. Raccoons were seen frequently on the farm. 

It was suspected that this straw bedding had been 

contaminated by raccoons, which had frequently 

been seen on the farm.

Kwiecen and others 

(1993).

Domestic sheep Idaho Ranch 1 A 3-day-old lamb had been moribund since birth and 

had Baylisascaris NLM. The pregnant ewe must 

have ingested infective eggs, and larvae migrated 

transplacentally into the developing lamb.

Anderson (1999).
Appendix 3 119

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.—Continued

[CNS, central nervous system; NR, not reported; >, greater than]

Animal(s) Location Facility type

Number of 

deaths or 

animals 

affected

Source of infection Reference

Ring-necked 

pheasants

Wisconsin Commercial 

ranch

65 Chronic, insidious losses from neurological disease 

over 2 years were observed at a ranch where 

200–400 pheasants were raised. The problem 

started when contaminated straw bedding was 

obtained from a neighbor’s barn.

Kazacos and others 

(1986).

Nutria Germany Breeding 

farm

65 Affected young nutria were temporarily kept in a 

pen that previously housed raccoons.

Koch and Rapp 

(1981).

Canaries California Private 

aviary

12 Raccoons were commonly seen around the aviary 

holding 22 birds.

Loretti and others 

(2008).

Pigeons Oregon Private 

collection

10 The source of infection in a collection of 60 birds 

was not identified. 

Helfer and Dickinson (1976).

Chukar Washington, 

D.C.

Private 

collection

1 An isolated case involved a collection of 30 birds. 

The source of infection was not determined, 

although raccoons and skunks were numerous in 

the area.

Sass and Gorgacz 

(1978).

Red (silver) 

foxes

Iowa Private 

collection

4 Four affected foxes were housed in a cage previously used for raccoons; five others housed in a cage 

previously used for rabbits remained normal.

Larson and Greve 

(1983).

Ostrich and 

emus

Indiana Private 

zoological 

collection

3 One ostrich and two emus were housed in a dirtfloored pen. Raccoons were housed in a loft 

directly overhead; feces from their cage dropped 

into the ostrich-emu pen below. Infective B. procyonis eggs were found in soil samples from the 

pen of the birds.

Kazacos and others 

(1991).

North American 

porcupines

New Jersey Rehabilitation 

facility

3 Three porcupines that died were housed in pens 

previously used for raccoons. A porcupine housed 

separately remained healthy.

Medway and others 

(1989).

Guinea pigs Missouri Research 

facility

30 Thirty of 50 animals in an accredited laboratory 

animal facility died with NLM. Bedding that was 

stored in an outdoor bin accessible by raccoons 

was unwittingly brought into the facility. Raccoon 

feces were found in the storage site and in bedding.

Van Andel and others (1995).

Guinea pigs Nova Scotia Research 

facility

2 Accidental cross-contamination was suspected from 

adult raccoons housed on the same premises.

Craig and others 

(1995).

Rhesus 

macaques

Southeastern 

United 

States

Research 

facility

13 Twenty-one macaques were from an outdoor breeding facility, where raccoons were common. Infections without clinical signs were discovered as 

part of another study.

Gozalo and others 

(2008).

Prairie dogs New York Research 

facility

3 Fifty-two prairie dogs at a medical research facility 

were obtained from a wildlife farm in Wisconsin 

where they had been temporarily housed in pens 

previously used for raccoons.

Dixon and others 

(1988).

Rabbits Japan Wildlife park 36 Rabbits housed in dirt-floored enclosures developed 

neurological signs after flooding from a typhoon. 

Raccoon feces and soil from raccoon pens or 

cages were dispersed by floodwaters distributing 

B. procyonis eggs into the rabbit enclosures.

Sato and others 

(2002, 2003).
120 Baylisascaris Larva Migrans

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.—Continued

[CNS, central nervous system; NR, not reported; >, greater than]

Animal(s) Location Facility type

Number of 

deaths or 

animals 

affected

Source of infection Reference

Japanese 

macaques

Japan Zoo 9 A colony of about 30 macaques was housed in a 

safari-style zoo in an open living space shared 

with black bears that were infected with Baylisascaris transfuga. A raccoon pen was adjacent, and 

feces drained directly into the macaque pen. B. 

procyonis infection was suspected based on severe 

damage to the brain and scarcity of larvae in the 

viscera of affected macaques.

Sato and others 

(2005).

Beaver Dublin, 

Ireland

Zoo >3 A zoo was unable to raise beavers to adulthood due 

to neurologic disease from NLM. The source of 

ascarid larvae in the animals was undetermined, 

but morphologically they were Baylisascaris spp.

Kelly and Innes 

(1966).

Nutria Michigan Zoo 20 Thirty-five nutria were fed tree branches as a dietary 

supplement and bedded on straw. The branches 

were collected from a wooded area where 

raccoons and skunks lived, and the onset and 

cessation of problems followed the initiation and 

cessation of feeding the tree branches. However, 

contaminated straw bedding could also have been 

the source.

Dade and others 

(1977).

Marmosets and 

tamarins

Texas Zoo 5 Skunks were also housed in the marmoset enclosures, and lived on the ground. Ascarid eggs 

(B. columnaris) were found in the skunk feces.

Huntress and 

Spraker (1985); 

K.R. Kazacos 

and P.L. Wolff, 

unpub. data 

(1986).

Blue-and-yellow 

and scarlet 

macaws

Nebraska Zoo 8 Ten macaws were placed in an island exhibit that 

was accessible to raccoons. Baylisascaris eggs 

were found in fecal and soil samples from the 

island, including in the elevated macaw food pans. 

Trapped raccoons were positive for B. procyonis.

Armstrong and 

others (1987, 

1989).

Red 

kangaroos

Michigan Zoo 11 Wild raccoons were common in the zoo. Raccoon 

feces were found in the pens housing 20 red kangaroos and on overhead boardwalks that traversed 

through the zoo.

Agnew and others 

(1994).

Emus Kansas Zoo 2 Four emus were housed in an indoor-outdoor pen. 

Raccoons were common on zoo grounds and had 

access to the outdoor pen. Also, straw bedding 

used by the original source of the emus may have 

been contaminated.

Suedmeyer and 

others (1996).

Black-and-white 

ruffed lemurs 

and emus

Oklahoma Zoo 5 Two lemurs and three emus were housed in an outdoor wire pen. Contamination of the enclosure by 

wild raccoons that were frequently seen in the zoo 

was suspected, and 57 raccoons were trapped near 

the enclosure.

Campbell and 

others (1997).
Appendix 3 121

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.—Continued

[CNS, central nervous system; NR, not reported; >, greater than]

Animal(s) Location Facility type

Number of 

deaths or 

animals 

affected

Source of infection Reference

Golden-headed 

lion 

tamarins 

Maryland 

and 

California

Zoos 3 At two zoos, tamarins were housed in outdoor pens 

with overhanging foliage; one zoo housed two 

tamarins and the other housed three. Raccoons 

and skunks were persistent problems within the 

zoos. Both tamarins at one zoo and one at the 

other died, and the other two remained healthy.

Pessier and others 

(1997); 

Stringfield 

and Sedgwick 

(1997).

Indian fruit bat, 

yellow-footed 

rock wallabies, lion 

tamarins, redtailed black 

cockatoo, 

thick-billed 

parrot, galah

California Zoo >15 Raccoons and skunks had free-run of the zoo, with 

widespread contamination. Roundhouse exhibits 

with wire tops allowed feces to fall into pens. 

Large wire mesh and overhanging foliage allowed 

animal access to the zoo and exhibits. A major 

control effort reduced the problem.

Stringfield and 

Sedgwick 

(1997).

White-handed 

gibbon

Kansas Zoo 1 A gibbon developed CNS signs after being placed 

in an island exhibit in which wild raccoons had 

established latrines. The gibbon spent time under 

its shelter, where raccoon feces with infective 

Baylisascaris eggs were found.

Ball and others 

(1988).

Orangutan Wisconsin Zoo 1 A 32.5-year-old orangutan developed worsening 

CNS disease and was euthanized. It had long-term 

exposure to an enclosure with raccoon latrines 

and had exhibited geophagia.

Hanley and others 

(2006).

Titi monkey Maryland Zoo 2 Two titi monkeys developed progressive CNS 

disease, and one responded to treatment. Raccoon 

latrines were found near their indoor and outdoor 

display enclosures.

Beck and others 

(2010).

Brushturkey Indiana and 

Missouri

Zoo 1 Wild raccoons were a common nuisance around the 

rearing pens, and contamination by hose-washing 

around the pens was suspected as the source of 

infection.

Kazacos and others 

(1982).

Burrowing 

parrots, 

crested 

screamers, 

porcupine

Manitoba Zoo 7 The animals were housed in chainlink mesh 

enclosures; raccoons had access to the tops of 

the enclosures and were observed on them. One 

porcupine, two screamers, and four of five parrots 

died.

Thompson and others (2008).

Amazon and 

Cuban parrots, 

rosy-faced 

lovebirds, 

rainbow and 

Swainson’s 

lorikeets

Ontario

 (Toronto)

Zoo 29 Raccoons and raccoon fecal contamination were 

both common on zoo grounds, especially around 

the aviary. Latrines were found on the tops of wire 

enclosures.

Russell and others 

(2005); Russell 

(2006).
122 Baylisascaris Larva Migrans

Appendix 3. Cases of Baylisascaris neural larval migrans in animal facilities.—Continued

[CNS, central nervous system; NR, not reported; >, greater than]

Animal(s) Location Facility type

Number of 

deaths or 

animals 

affected

Source of infection Reference

De Brazza’s 

monkeys

Indiana Zoo 2 The monkeys had subtle CNS disease following 

exposure to a contaminated roundhouse enclosure. 

Raccoon latrines were found on the top of the 

enclosure.

C.L. Eng and 

K.R. Kazacos, 

unpub. data 

(1997).

Mouse lemur California Zoo 1 Exposure to contaminated tree limbs and branches 

put into the exhibit was suspected.

K.R. Kazacos 

and F.H. Dunker, 

unpub. data 

(1995).

Patagonian 

maras and 

capybara

Illinois Zoo 5 All animals were exposed to raccoon fecal contamination in their enclosures from wild raccoons on 

zoo grounds; four maras and one capybara died.

K.R. Kazacos and 

others, unpub. 

data (1987 

[capybara] and 

1989 [maras]).
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/
Kazacos—Baylisascaris Larva Migrans—Circular 1412

ISSN 1067-048X (print)

ISSN 2330-5703 (online)

http://dx.doi.org/10.3133/cir1412