U.S. Department of the Interior

U.S. Geological Survey

Scientific Investigations Report 2012–5271

U.S. Geological Survey, National Wildlife Health Center, 

2011 Report of Selected Wildlife Diseases
Cover images, clockwise, starting top left:

David Green, National Wildlife Health Center, with blackbird (Photo by Allison Klein) 

NWHC staff and partners with cormorant (Photo by C. LeAnn White) 

Box turtle (Photo by Cindy Cunningham)

Bald eagle (Photo by J. Chris Franson) 

Background photo of snow geese (Photo by Bob Dusek)
U.S. Geological Survey, National Wildlife 

Health Center, 2011 Report of Selected 

Wildlife Diseases 

By David Green, Megan Hines, Robin Russell, and Jonathan Sleeman

Scientific Investigations Report 2012–5271

U.S. Department of the Interior

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

KEN SALAZAR, Secretary

U.S. Geological Survey

Marcia K. McNutt, Director

U.S. Geological Survey, Reston, Virginia: 2012

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Suggested citation:

Green, D., Hines, M., Russell, R., and Sleeman, J., 2012, U.S. Geological Survey, National Wildlife Health Center , 

2011 report of selected wildlife diseases: U.S. Geological Survey Scientific Investigations Report 2012–5271, 32 p. 

plus 1 app.
iii

Acknowledgements

We thank our many partners and submitters for their vigilance in monitoring for wildlife disease 

outbreak events and for submitting cases to us for diagnostic evaluation. 

Contents

Conversion Factors .......................................................................................................................................v

Introduction.....................................................................................................................................................1

Materials and Methods.................................................................................................................................1

Selected Wildlife Diseases ..........................................................................................................................3

Avian Diseases......................................................................................................................................3

Botulism Type C............................................................................................................................3

Cause or Etiology ................................................................................................................3

Host Species and Annual Distribution ............................................................................3

Geographic Distribution.....................................................................................................3

Seasonality ..........................................................................................................................3

Signs and Lesions...............................................................................................................4

Significance and Zoonotic Risk........................................................................................5

Selected References..........................................................................................................5

Botulism Type E............................................................................................................................6

Cause or Etiology ................................................................................................................6

Host Species and Annual Distribution ............................................................................6

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

Seasonality ..........................................................................................................................6

Signs and Lesions...............................................................................................................6

Significance and Zoonotic Risk........................................................................................8

Selected References..........................................................................................................8

Avian Cholera ..............................................................................................................................9

Cause or Etiology ................................................................................................................9

Host Species........................................................................................................................9

Geographic Distribution.....................................................................................................9

Seasonality ........................................................................................................................11

Signs and Lesions.............................................................................................................11

Significance and Zoonotic Risk......................................................................................11

Selected References........................................................................................................11
iv

Salmonellosis..............................................................................................................................12

Cause or Etiology ..............................................................................................................12

Host Species and Annual Distribution ..........................................................................12

Geographic Distribution...................................................................................................12

Seasonality .......................................................................................................................12

Signs and Lesions.............................................................................................................12

Significance and Zoonotic Risk......................................................................................14

Selected References........................................................................................................14

Exotic Trematode Infections ....................................................................................................15

Cause or Etiology ..............................................................................................................15

Host Species......................................................................................................................15

Geographic Distribution...................................................................................................15

Seasonality ........................................................................................................................16

Signs and Lesions.............................................................................................................17

Significance and Zoonotic Risk......................................................................................17

Selected References........................................................................................................17

Lead Poisoning (Plumbism)......................................................................................................18

Cause or Etiology ..............................................................................................................18

Host Species and Annual Distribution ..........................................................................18

Geographic Distribution...................................................................................................18

Seasonality .......................................................................................................................18

Signs and Lesions.............................................................................................................20

Significance and Zoonotic Risk......................................................................................20

Selected References........................................................................................................20

Virulent Newcastle Disease Virus ..........................................................................................21

Cause or Etiology ..............................................................................................................21

Host Species and Annual Distribution ..........................................................................21

Geographic Distribution...................................................................................................21

Seasonality .......................................................................................................................22

Signs and Lesions.............................................................................................................22

Significance and Zoonotic Risk......................................................................................23

Selected References........................................................................................................23

West Nile Virus...........................................................................................................................24

Cause or Etiology ..............................................................................................................24

Host Species and Annual Distribution ..........................................................................24

Geographic Distribution...................................................................................................24

Seasonality ........................................................................................................................24

Signs and Lesions.............................................................................................................25

Significance and Zoonotic Risk......................................................................................25

Selected References........................................................................................................26
v

Selected Herpetological Diseases ..................................................................................................27

Amphibian Chytridiomycosis ...................................................................................................27

Cause or Etiology ..............................................................................................................27

Host Species and Annual Distribution ..........................................................................27

Geographic Distribution...................................................................................................27

Seasonality .......................................................................................................................28

Signs and Lesions.............................................................................................................29

Significance and Zoonotic Risk......................................................................................29

Selected References........................................................................................................29

Ranavirus.....................................................................................................................................30

Cause or Etiology ..............................................................................................................30

Host Species and Annual Distribution ..........................................................................30

Geographic Distribution...................................................................................................30

Seasonality .......................................................................................................................30

Signs and Lesions.............................................................................................................32

Significance and Zoonotic Risk......................................................................................32

Selected References........................................................................................................32

Appendix 1.....................................................................................................................................................33

Figures

1. Graph showing occurrence of botulism type C in wild birds 2001–10, by order................3

2. Maps showing prevalence of botulism C diagnosed in wild bird disease 

outbreaks, as a percentage of all wild bird disease outbreaks, by State, 

for A, 2001–5 and B, 2006–10.......................................................................................................4

3. Graph showing monthly number of cases of botulism type C in wild birds........................5

4. Graph showing annual number of cases of botulism type C in wild birds..........................5

5. Graph showing occurrence of botulism type E in wild birds 2001–10, by order................6

6. Maps showing prevalence of botulism E diagnosed in wild bird disease 

outbreaks, as a percentage of all wild bird disease outbreaks, by State, 

for A, 2001–5 and B, 2006–10.......................................................................................................7

7. Graph showing annual number of cases of botulism type E in wild birds ..........................8

8. Graph showing monthly number of cases of botulism type E in wild birds ........................8

9. Graph showing occurrence of avian cholera in wild birds, 2001–10, by order..................9

10. Maps showing prevalence of avian cholera diagnosed in wild bird disease 

outbreaks, as a percentage of all wild bird disease outbreaks, by State, 

for A, 2001–5 and B, 2006–10.....................................................................................................10

11. Graph showing annual number cases of avian cholera in wild birds ...............................11

12. Graph showing monthly number cases of avian cholera in wild birds .............................11

13. Graph showing occurrence of salmonellosis in wild birds 2001–10, by order.................12

14. Maps showing prevalence of salmonellosis diagnosed in wild bird disease 

outbreaks, as a percentage of all wild bird disease outbreaks, by State, 

for A, 2001–5 and B, 2006–10.....................................................................................................13

15. Graph showing annual number of cases of salmonellosis in wild birds...........................14

16. Graph showing monthly number of cases of salmonellosis in wild birds.........................14

17. Graph showing occurrence of exotic trematode infections in wild birds 2001–10, 

by order ........................................................................................................................................15
vi

 18. Maps showing prevalence of exotic trematode infections diagnosed in wild bird 

Multiply By To obtain

Length

meter (m) 3.281 foot (ft) 

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

°F=(1.8×°C)+32
U.S. Geological Survey, National Wildlife Health Center, 

2011 Annual Report of Selected Wildlife Diseases

By David Green1, Megan Hines2, Robin Russell1, and Jonathan Sleeman1

Introduction

The National Wildlife Health Center (NWHC) was 

founded in 1975 to provide technical assistance in identifying, 

controlling, and preventing wildlife losses from diseases, conduct research to understand the impact of diseases on wildlife 

populations, and devise methods to more effectively manage 

these disease threats. The impetus behind the creation of the 

NWHC was, in part, the catastrophic loss of tens of thousands 

of waterfowl as a result of an outbreak of duck plague at the 

Lake Andes National Wildlife Refuge in South Dakota during 

January 1973. In 1996, the NWHC, along with other Department of Interior research functions, was transferred from the 

U.S. Fish and Wildlife Service to the U.S. Geological Survey 

(USGS), where we remain one of many entities that provide 

the independent science that forms the bases of the sound 

management of the Nation’s natural resources. Our mission is 

to provide national leadership to safeguard wildlife and ecosystem health through dynamic partnerships and exceptional 

science.

The main campus of the NWHC is located in Madison, 

Wis., where we maintain biological safety level 3 (BSL–3) 

diagnostic and research facilities purposefully designed for 

work with wildlife species. The NWHC provides research and 

technical assistance on wildlife health issues to State, Federal, 

and international agencies. In addition, since 1992 we have 

maintained a field station in Hawaii, the Honolulu Field Station, which focuses on marine and terrestrial natural resources 

throughout the Pacific region.

The NWHC conducts diagnostic investigations of 

unusual wildlife morbidity and mortality events nationwide 

to detect the presence of wildlife pathogens and determine the 

cause of death. This is also an important activity for detecting 

new, emerging and resurging diseases. The NWHC provides 

this crucial information on the presence of wildlife diseases 

to wildlife managers to support sound management decisions. 

The data and information generated also allows for further indepth analyses for determining the biological and ecological 

1

U.S. Geological Survey 

2

School of Veterinary Medicine, University of Wisconsin

significance of disease events, detecting disease trends over 

time and space, as well as detecting any significant changes to 

how diseases manifest in the field. Moreover, this information 

allows us to gain insight into the significance of future wildlife 

disease events.

The purpose of this report is to provide a sample of 

NWHC data that are available from our Laboratory Information Management System (LIMS). These data are presented in 

summary format with minimal statistical analysis and interpretation. The goal is to share these data with wildlife managers and other stakeholders, promote the use of NHWC data, 

and encourage the sharing of wildlife disease data to improve 

temporal and geographic surveillance coverage. Continued 

national surveillance for wildlife diseases is essential for 

providing early detection and warning of events that have the 

potential to result in harm to human health, economic losses, 

declines in wildlife populations, and subsequent ecological 

disturbances. Increased collaboration, coordination, and sharing of surveillance data will enhance this Nation’s ability to 

detect and respond to wildlife disease threats. 

Materials and Methods

Biological samples, mostly carcasses, are submitted voluntarily to the NWHC for diagnostic evaluation to determine 

the cause of death. These carcasses and samples constitute the 

basis of the data used in this report. After a group of carcasses 

or “submission” arrives at the NWHC, each submission is 

given a case number, and then each carcass within the submission is given an accession number. Multiple carcasses from 

a disease event are often submitted for diagnosis; therefore, 

each case number is usually associated with more than one 

accession number. These numbers—along with information 

regarding the species, geographic location where the carcass 

was found, the submitting agency, and the dates of collection and submission—are entered into the LIMS database 

at the NWHC. The diagnoses presented in this report were 
2 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

determined by NWHC pathologists using gross and histological evaluations and ancillary diagnostic tests as needed, and 

were based on published or established diagnostic criteria, 

including World Organization for Animal Health (OIE) published criteria where appropriate. 

For the summaries contained within this report, we present data for diseases that are on the World Organization for 

Animal Health (OIE) list of reportable diseases or among the 

top 10 most common diagnoses at NWHC, or both. Background information about and data for eight avian diseases are 

included in the report: botulism C, botulism E, avian cholera, 

salmonellosis, exotic trematode infections, lead poisoning 

(plumbism), virulent Newcastle disease virus, and West Nile 

virus. Data for two diseases of amphibians and reptiles (herpetological diseases), chytridiomycosis and ranavirus infection, also are included. The number of cases of each disease is 

summarized in multiple ways: (1) the number of cases of each 

disease reported each year (the annual count); (2) the number 

of cases of each disease reported each month (the monthly 

count); (3) the number of cases as proportion of total cases for 

all diseases (either avian or herpetological diseases); (4) the 

number of cases as proportion of total cases for all diseases 

in a State; and (5) the proportion of cases for each disease by 

taxonomic grouping.

Annual counts were calculated by summing the total 

number of cases in a year for each year from 2001 to 2010, 

and monthly counts were calculated by summing the total 

number of cases in each month pooled across years. For avian 

diseases, the number of cases as a proportion of total cases 

was calculated by summing the total number of avian cases for 

each of the eight diseases included in this report for each year, 

and then dividing the total number of cases for a particular 

avian disease by the total for all avian diseases. The equivalent 

was done for the herpetological diseases, ranavirus and chytridiomycosis. For the proportion by State, the total number of 

cases in each state was summed separately for avian diseases 

(the eight included in the report) and herpetological diseases 

(ranavirus and chytridiomycosis), and then the number of 

cases for each disease was divided by the total number of 

avian or herpetological cases in the State. 

Lastly, we report the proportion of cases of an individual 

disease that included at least one member of a particular 

taxonomic group. For avian diseases we have grouped species 

by taxonomic order: Anseriformes (ducks, geese, swans), 

Charadriiformes (shorebirds, gulls, terns), Ciconiiformes (herons, egrets), Columbiformes (doves, pigeons), Falconiformes 

(hawks, eagles, falcons), Galliformes (upland game birds), 

Gaviiformes (loons), Passeriformes (songirds), Pelecaniformes 

(pelicans, cormorants), Gruiformes (cranes, rails, coots), Podicipediformes (grebes), and Strigiformes (owls). For herpetological diseases we categorized specimens as frog, tree frog, 

toad, salamander and newt, and turtle. Each case could be 

included more than once in this summary, if the case contained 

specimens from more than one taxonomic group. In other 

words, the sum of the proportions of cases in each taxonomic 

group for one disease could exceed 100 percent.

Voluntary case submissions constitute a passive sampling technique, and the NWHC has no quantitative measure 

of survey effort. Therefore, we exercise caution in interpreting fluctuations in the number of cases submitted as an index 

of temporal or spatial trends in the incidence of a disease. 

Many unmeasured factors—changes in the number of field 

personnel or the amount of funding, changes in the detection 

probability of the carcasses (increased or decreased predation, 

for example), and the acquisition or loss of local diagnostic 

experts—can result in a change in submission rates to the 

NWHC. Furthermore, additional cases were likely submitted 

to State or regional diagnostic laboratories for evaluation and 

would not be included in this report. Thus, we report raw data 

with minimal interpretation to provide information on where, 

when, and in what species particular diseases have been 

detected in NWHC submissions. We believe these data, combined with other sources of information, may generate testable 

hypotheses regarding disease status and trends and provide a 

launching point toward the better understanding of the ecology 

of wildlife diseases. The data used in this report can be found 

in the appendix.
Selected Wildlife Diseases 3

Selected Wildlife Diseases

Avian Diseases

Botulism Type C

Cause or Etiology

Botulism type C is an intoxication by a natural toxin 

produced by the bacterium, Clostridium botulinum. This 

anaerobic bacterium produces multiple types of botulinum 

toxins that are referred to as types A through G. To date, only 

botulinum toxins types C and E are known to cause morbidity 

and mass mortality events in wild birds in the United States. 

In birds, botulism type C is caused by ingestion of the toxin 

from the environment, and is usually associated with ingestion 

of invertebrates in and adjacent to rotting piles of vegetation 

(usually in or adjacent to ponds, lakes, or shallow wetlands) or 

by ingestion of one or more maggots from rotting carcasses. 

Unlike cases of botulism in humans and livestock, cases of 

botulism associated with anaerobic growth of the bacterium in 

the intestine or wounds are not documented or are very rare in 

birds.

Host Species and Annual Distribution

Botulism type C affects and kills a much greater 

taxonomic range of birds that botulism type E. Waterfowl 

such as ducks, geese, and swans most often are involved 

(fig. 1), but gulls, terns, pelicans, egrets, ibises, herons, stilts, 

coots, sandpipers, sanderlings, and plovers also have been 

test-positive for botulism type C. Infrequently, botulism is 

confirmed in yellowlegs, godwits, avocets, and turnstones. A 

diagnosis of botulism of any type is difficult in small shore 

birds and most passerines, because of their small body size 

and the difficulty in collecting a volume of blood sufficient 

for the diagnostic test. Often, blood samples are pooled from 

multiple small birds of the same species from the same event 

for a single test. The taxa of birds in which botulism has been 

observed prior to 2001, but was not confirmed in 2001–10, are 

raptors, owls, pigeons, doves, quail, pheasants, grouse, and 

most songbirds.

Geographic Distribution

Die-offs due to botulism type C between 2001 and 

2010 were documented in 29 States, Puerto Rico, and the 

Pacific Islands of Laysan and Palau (fig. 2). Mortality events 

due to botulism type C have involved endangered species, 

including Hawai’ian stilts (black-necked stilt or Ae’o; 

Himantopus mexicanus knudseni), Laysan ducks (Laysan 

teal; Anas laysanensis), Hawai’ian coot (‘Alae Ke’oke’o; 

Fulica americana alai), Hawai’ian duck (Koloa Maoli; 

Anas wyvilliana) and others. Distribution of botulism type 

C remains similar to previously reported cases (Rocke and 

Friend, 1999).

Seasonality

Botulism type C has a clear 

seasonality with most cases occurring 

during summer months and peaking 

in August in temperate regions. In 

southern States, cases may occur every 

month of the year, but rarely at times 

when snow covers the ground or ice 

covers wetlands. The onset of cases is 

usually in late June, and the number of 

mortality events increases steadily in 

July and August (fig. 3). The number of 

annual mortality events peaked in 2006 

(fig. 4) at 49 cases and has declined 

steadily since, but the number of cases 

increased slightly in 2009. 

Proportion of diagnostic cases

Taxomonic order

0.0

0.2

0.4

0.6

0.8

1.0

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 1. Occurrence of botulism type C in wild birds, by order.

Figure 1. Occurrence of botulism type C in wild birds 2001–10, by order.
4 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

EXPLANATION

Botulism C, as a percentage 

of total avian cases 

investigated at NWHC

0

1 to 15

16 to 30

31 to 50

51 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 2. Maps showing the prevalence of botulism C diagnosed in wild bird disease outbreaks, as a 

percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Signs and Lesions

Field observations 

usually begin with the 

appearance of numerous 

carcasses along a 

shoreline and with fewer 

sick birds. The signs of 

botulism types C and E 

are indistinguishable, 

and they include 

lethargy, reluctance to 

fly or swim, limb and 

neck paralysis, and 

paralysis of the third 

eyelids. Birds found 

dead on open water may 

have had neck paralysis 

sufficient to prevent 

the head from being 

immersed in water; these 

birds may have internal 

abnormalities consistent 

with drowning. Botulism 

typically is lethal to 

birds within hours; so 

many birds are in fair 

to good body condition 

and without obvious 

internal abnormalities. 

Occasionally, ingested 

maggots may be 

found in the lumen of 

the distal esophagus, 

proventriculus, or 

gizzard; ingested 

maggots, when tested, 

usually are heavily laden 

with botulinum toxin(s). 

Figure 2. Prevalence of botulism C diagnosed in wild bird disease outbreaks, as a percentage of all 

wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
Selected Wildlife Diseases 5

0

20

40

60

80

100

Month

Number of diagnostic cases

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Monthly number of cases of botulism type C in wild bird carcasses.

Figure 3. Monthly number of cases of botulism type C in wild 

birds.

0

10

20

30

40

50

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Annual number of cases of botulism type C in wild birds.

Figure 4. Annual number of cases of botulism type C in wild 

birds.

Significance and Zoonotic Risk

Botulism type C may affect a variety of wildlife 

and domestic animals, but type C is much less 

hazardous to humans than type E. Although animals 

that prey on sick birds with botulism or scavenge 

on their carcasses may contract secondary botulism, 

this is rarely documented; however, in 2006—the 

year of the peak number of botulism type C cases 

(49)—three dogs that were trained and working as 

retrievers of waterfowl were diagnosed with botulism 

type C. One dog was used by a biologist to retrieve 

carcasses of waterfowl during confirmed die-offs due 

to botulism type C, and another dog was used as a 

retriever by duck hunters at a site where botulism had 

occurred. All three dogs became partially paralyzed 

and then recovered after a week of intensive veterinary 

clinical care. Because carcasses may provide suitable 

anaerobic growth media for Clostridium botulinum

and become infested with toxin-laden maggots, 

these maggot-infested carcasses become the source 

of additional cases of botulism when birds feed on 

maggots. Smaller invertebrates that feed on botulinumladen carcasses may be the source of botulism 

for shore birds such as plovers, sanderlings, and 

sandpipers.

Selected References

Rocke, T.E., and Bollinger, T.K., 2007, Avian botulism, chap. 21 of Thomas, N.J., Hunter, D.B., 

and Atkinson, C.T., eds., Infectious diseases of 

wild birds: Ames, Iowa, Blackwell Publishing, p. 

377–416.

Rocke, T.E., and Friend, M., 2009, Avian botulism, 

chap. 38 of Friend, M., and Franson, J.C., eds., Field 

manual of wildlife diseases: general field procedures and diseases of birds: U.S. Geological Survey 

Information and Technology Report 1999–001, p. 

271–281.
6 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

Botulism Type E

Cause or Etiology

Botulism type E is intoxication by a natural toxin produced by the bacterium, Clostridium botulinum. This anaerobic bacterium produces multiple types of botulinum toxins that 

are referred to as types A through G. To date, only botulinum 

toxins types C and E are known to cause morbidity and 

massive mortality events in wild birds in the United States. 

Botulinum toxins A, B, E, and, rarely, F are usually implicated 

in human cases of botulism. In birds, the disease, botulism, 

is caused by ingestion of the toxin from the environment and 

usually is associated with ingestion of fish with botulism, 

ingestion of invertebrates in and adjacent to rotting piles of 

vegetation (usually in or adjacent to ponds, lakes, or shallow wetlands), or by ingestion of one or more maggots from 

rotting carcasses. Unlike cases of botulism in humans and livestock, cases of botulism associated with anaerobic growth of 

the bacterium in the intestine or wounds are not documented 

or are very rare in birds.

Host Species and Annual Distribution

Botulism type E affects many more fish-eating species 

of birds than botulism type C (fig. 5). During the period of 

2001–5, this disease was reported predominantly along the 

shorelines of the Great Lakes (fig. 6A), and 8 of 10 mortality 

events involved loons, cormorants, pelicans, or gulls. A single 

common loon (Gavia immer) from Maine was detected with 

Figure 5. Occurrence of botulism type E ...

Proportion of diagnostic cases

0.0

0.2

0.4

0.6

0.8

1.0

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 5. Occurrence of botulism type E in wild birds 2001–10, by order.

botulism type E in 2005. American coots (Fulica americana) 

and multiple species of ducks were affected in 2 of 10 other 

mortality events. During 2006–10, mortality events due to 

botulism type E were an annual occurrence on the shores of 

the Great Lakes, but additional cases were detected in the 

Western States of North Dakota, Montana, and Idaho (fig. 6B). 

At present, it is unclear whether botulism type E is spreading westward or simply was not detected in the three Western 

States until 2009 and 2010. In the years 2006–10, affected 

species again were mostly fish-eating birds; and grebes, terns, 

sandpipers, and plovers were affected in significant numbers in 

addition to the loons, cormorants, pelicans, and gulls affected 

during 2001–5. Seven mortality events during 2001–5 and 18 

during 2006–10 were attributed to botulism type E. It is not 

clear whether increasing events are due to increased awareness 

and surveillance for this disease or other unknown factor(s). 

The seasonality and possible host cycles of botulism type E 

are being studied extensively.

 Geographic Distribution

Botulism type E occurs predominantly in States and 

Canadian Provinces that border the five Great Lakes (fig. 6A). 

Only five mortality events have occurred since 2005 in States 

not bordering the Great Lakes: Maine, North Dakota with two 

mortality events, Montana, and Idaho (fig. 6B). 

Seasonality

The vast majority of mortality events due to botulism 

type E occur between June and November annually (fig. 7). 

Mortality events tend to peak yearly in 

the months of July and October. Cases 

of all types of botulism peak during 

the summer and autumn months and 

are infrequent during winter and spring 

months (fig. 8). 

Signs and Lesions

Field signs of botulism type E are 

indistinguishable from those of botulism type C (see the previous section on 

botulism type C), but field signs usually 

involve the appearance of dead fisheating birds along lake shores. Many of 

these birds are presumed to have ingested 

fish affected by botulism in open water, 

and after death, the carcasses wash 

ashore. The contents of the esophagus, 

crop, proventriculus, and ventriculus 

(gizzard) generally are not helpful in 

making a presumptive diagnosis of botulism, because, at present, it is not clear 

which species of fish may be more or less 

resistant to botulism. 
Selected Wildlife Diseases 7

EXPLANATION

Botulism E, as a percentage 

of total avian cases 

investigated at NWHC

0

1 to 10

11 to 25

26 to 50

51 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10

Figure 6. Maps showing the prevalence of botulism E diagnosed in wild bird disease outbreaks, as 

a percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 6. Prevalence of botulism E diagnosed in wild bird disease outbreaks, as a percentage of all 

wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
8 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

0

10

20

30

40

50

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Annual number of cases of botulism type E in wild birds.

Figure 7. Annual number of cases of botulism type E in wild birds.

0

20

40

60

80

100

Month

Number of diagnostic cases

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Figure 8. Monthly number of cases of botulism type E in wild birds.

Significance and Zoonotic Risk

Botulism type E is an important zoonosis, and immediately following confirmation of the disease (by mouse protection tests) it is a reportable disease to the Centers for Disease 

Control (CDC) Select Agent Program. This requirement also 

applies to Botulism Type C. Humans may be sickened or 

killed by botulism type E, so blood, tissues, and carcasses of 

birds suspected to have type E poisoning should be handled 

with personal protection equipment (“PPE”; that is, gloves, 

particulate masks, or respirators, and so forth). Domestic 

animals, including dogs, are susceptible to botulism, and 

there are cases of dogs that retrieve carcasses of dead birds 

subsequently developing botulism. Thus, dogs should not be 

used when collecting sick and dead birds suspected of having 

botulism. Management of disease outbreaks involves daily or 

twice daily collection of fish and bird carcasses to interrupt the 

carcass-maggot (or carcass-invertebrate) cycle of botulism. 

Selected References

Lafrancois, B.M., Riley, S.C., Blehert, D.S., and Ballmann, 

A.E., 2011, Links between type E botulism outbreaks, lake 

levels and surface water temperatures in Lake Michigan, 

1963–2008: Journal of Great Lakes Research v. 37, 

p. 86–91. 

Perez-Fuentetaja, A., Clapsadl, M.D., Getchell, R.G., Bowser, 

P.R., and Lee, W.T., 2011, Clostridium botulinum type E 

in Lake Erie: Inter-annual difference and role of benthic 

invertebrates: Journal of Great Lakes Research, v. 37, no. 2, 

p. 238–244, http://dx.doi.org/10.1016/jglr.2011.03.013.
Selected Wildlife Diseases 9

Avian Cholera 

Cause or Etiology

Avian cholera (avian pasteurellosis, fowl cholera) is a 

contagious infectious disease of many species of birds caused 

by the bacterium Pasteurella multocida. Avian cholera is not 

related to the bacterium and toxins that cause human cholera. 

Domestic and wild birds can be infected by many serotypes 

of P. multocida, but the predominant serotype in wild birds in 

the U.S. is serotype 1. Serotypes 3 and 4 occur occasionally 

in massive mortality events on the East Coast, but they were 

not observed during the 10 years of this report. Serotypes of 

P. multocida isolated from infected birds in Hawai’i (data not 

shown on maps and figures) tend to have uncommon serotypes 

and mixtures of serotypes such as serotypes 8, 12, and others. 

The bacterium is probably transmitted when birds ingest the 

organism from contaminated aquatic environments during 

seasons when birds are crowded. The bacterium may also be 

transmitted bird-to-bird and by aerosol. Studies by NWHC 

scientists (Samuel and others, 2005) have provided evidence 

that up to 2 percent of normal-appearing lesser snow (Chen 

caerulescens) and Ross’s geese (Chen rossii) are carriers of 

pathogenic serotypes of avian cholera. Many mortality events 

due to avian cholera are associated with seasonal migratory 

appearance of these geese in the Mississippi, Central, and 

Pacific Flyways. 

Host Species

It is probable that most species of birds are susceptible 

to infection by Pasteurella multocida, but mortality events 

predominantly involve waterfowl and shorebirds (fig. 9). The 

vast majority of birds dying with avian cholera are ducks, 

geese, swans, grebes, and coots, with lesser numbers of 

gulls. Small mortality events involving only 1–3 birds during 

2001–10 occurred in bald eagles (Haliaeetus leucocephalus), 

a red-tailed hawk (Buteo jamaicensis), a long-eared owl (Asio 

otus), black-necked stilts (Himantopus mexicanus), killdeer 

(Charadrius vociferous), cormorants, common ravens (Corvus 

corax), and other passerines. 

Geographic Distribution

All but three mortality events in 2001–10 were in States 

west of the Mississippi River in the Mississippi, Central, and 

Pacific Flyways (figs. 10A and B). In 2008, avian cholera was 

documented for the first time in Arkansas in lesser snow and 

Ross’s geese and ducks; but avian cholera had been detected 

for decades in States north, south, and west of Arkansas. From 

2001 to 2010, avian cholera was not detected in five Western 

States (Washington, Nevada, New Mexico, Colorado, and 

Montana) where it was regularly documented in the years 

prior to this report, and it was not detected in three northern 

Atlantic Coastal states (Maine, Maryland, and Virginia). 

The vast majority of serotypes involved in 

western mortality events were type 1, but 

data for much of 2001–5 are incomplete 

and comparison to the second 5-year period 

(2006–10) is not possible (figs. 10A and B). 

Most avian cholera events were in freshwater and estuarine environments, but several 

States of the Great Plains and Great Basin 

have numerous saline and hypersaline lakes 

(and ”potholes”) where mortality events 

due to avian cholera are frequent. Mortality 

events in three States east of the Mississippi 

River (Florida, Illinois, and Mississippi) 

involved lesser snow and Ross’s geese and a 

coot; it is suspected that populations of lesser 

snow and Ross’s geese are increasing, and 

overwintering populations may be expanding 

their winter ranges eastward. Data for avian 

cholera events in Hawai’i and Pacific Islands 

are incomplete.

Proportion of Diagnostic Cases

0.0

0.2

0.4

0.6

0.8

1.0

Figure 9. Graph showing the occurrence of avian cholera in wild bird carcasses, by family.

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes

Figure 9. Occurrence of avian cholera in wild birds, 2001–10, by order.
10 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

EXPLANATION

Avian cholera, as a percentage of total avian 

cases investigated 

at NWHC

0

1 to 10

11 to 25

26 to 50

51 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 10. Maps showing the prevalence of avian cholera diagnosed in wild bird disease outbreaks, 

as a percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 10. Prevalence of avian cholera diagnosed in wild bird disease outbreaks, as a percentage of 

all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
Selected Wildlife Diseases 11

Seasonality

Most mortality events due to avian cholera occur during winter months of November through March (fig. 12) on 

wintering grounds during periods of waterfowl and shorebird 

crowding. Cases of avian cholera occur in every month of the 

year, but casualty numbers during late spring, summer, and 

early autumn usually number in the dozens, although those 

during winter months involve hundreds or thousands of birds.

Signs and Lesions

Avian cholera is consistently a rapidly lethal systemic 

infection of wild birds. Most birds die in 8–48 hours following exposure. Consequently, most dead birds will be in fair to 

good body condition. Field and clinical signs in live birds are 

varied and attributed to bacterial septicemia. Signs include 

lethargy, weakness, reluctance to fly, walk, or swim, and 

occasionally, clear or muco-purulent nasal and oral discharges. 

Some birds may present with neurological signs. Sick birds 

often die within minutes of being collected or handled. At 

necropsy, the characteristic finding is minute white foci of 

necrosis in the liver (and occasionally the spleen), but white 

foci in the liver may occur with other infectious diseases such 

as salmonellosis and erysipelas. Another common abnormality 

is a thick, tenacious, yellow muco-purulent material within the 

small intestine. Petechial hemorrhages on the heart, liver, and 

other serosal surfaces may be present. 

Significance and Zoonotic Risk

Avian cholera is considered a low risk to humans because 

of differences in species susceptibility to the many serotypes 

of P. multocida. Birds and mammals may be infected by P. 

multocida through wounds from the bites of predators (especially cats); therefore, persons handling dead birds should 

wear gloves to prevent secondary infections of minor skin 

wounds and abrasions. When handling large numbers of carcasses and disposing of them by burying or burning, there is a 

significant possibility of aerosol transmission of the bacterium, 

especially in confined spaces; persons should wear masks 

(particle respirators) when handling numerous avian carcasses. 

Carcasses should be disposed by methods that make them 

unavailable to susceptible scavengers such as mice, crows, 

gulls, and eagles.

Selected References

Samuel, M.D., Botzler, R.G., and Wobeser, G.A., 2007, Avian 

cholera, in Thomas, N.J., Hunter, D.B., and Atkinson, 

C.T., eds., Infectious diseases of wild birds: Ames, Iowa, 

Blackwell Publishing, p. 239–269.

Samuel, M.D., Shadduck, D.J., Goldberg, D.R., and Johnson, 

W.P., 2005, Avian cholera in waterfowl: The role of Lesser 

Snow and Ross’s geese as disease carriers in the Playa 

Lakes Region: Journal of Wildlife Diseases, v. 41, no. 1, 

p. 48–57. 

birds.

birds.

0

5

10

15

20

25

30

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 11. Annual number cases of avian cholera in wild 

0

5

10

15

20

25

30

Number of diagnostic cases

Monthly number of cases of avian cholera in wild bird carcasses.

Month

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Figure 12. Monthly number cases of avian cholera in wild 
12 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

Salmonellosis

Cause or Etiology

Salmonella bacteria are a fairly common cause of systemic lethal infections in songbirds (passerines) in the U.S., 

and many cases are associated with specific backyard birdfeeding stations. The vast majority of cases of salmonellosis 

in passerine birds linked to backyard birdfeeders are due to 

Salmonella typhimurium. 

Host Species and Annual Distribution

About half of all cases of salmonellosis are in passerines 

associated with bird-feeding stations (fig. 13). Affected species include pine siskins (Carduelis pinus), common redpolls 

(Carduelis flammea), American goldfinches (Carduelis tristis), 

house sparrows (Passer domesticus), crossbills (Loxia spp.), 

grosbeaks, cowbirds, and various species of finches. Mortality 

events also occur regularly during nesting season in colonynesting birds such as common terns (Sterna hirundo), doublecrested cormorants (Phalacrocorax auritus), cattle egrets 

(Bubulcus ibis), herons, pelicans, and gulls. Infrequently, 

infections are detected in doves, hawks, and grebes. There 

were no cases of Salmonella pullorum in wild gamebirds 

(gallinaceous birds) from 2001 to 2010. Salmonella spp. are 

isolated frequently from only the intestine (colon or cloaca) of 

birds and reptiles, but these limited infections are not included 

with these systemic lethal infections. One trend may have 

emerged in 2001–10 for pelicans and cormorants. Salmonellosis in these colony-nesting birds was considered “infrequent” 

(Friend, 1999), but with increased monitoring of colonynesting birds for virulent Newcastle Disease virus (vNDV) 

and West Nile virus, salmonellosis must now be considered a 

common cause of mortality in these birds.

Geographic Distribution

Salmonellosis occurs nationwide (fig. 14) and is 

considered a global zoonosis.

Seasonality 

Salmonellosis has two trends: cases among songbirds 

during winter months that are associated with backyard feeding stations and cases among shorebirds such as gulls, cormorants, and terns at nesting sites during nesting season (fig. 16). 

The number of cases of salmonellosis in colony-nesting birds 

increases greatly during summer months. The number of cases 

during winter months in songbirds and in pigeons and doves 

(Columbiformes) that frequent garden birdfeeders probably is 

significantly underreported. 

Signs and Lesions

Passerine birds with salmonellosis usually present as 

lethargic and emaciated. Bacterial caseous granulomas are 

most commonly detected in the esophagus and crop; these 

granulomas often are large enough to be palpable in the live 

bird or intact carcass. Granulomas in passerines also are found 

in the intestine, cloaca, liver, and spleen. Granulomas in the 

liver and spleen often are pinpoint-sized yellow foci. Salmonella infections in shorebirds such as gulls, terns, cormorants, 

egrets, herons, and pelicans may present as acute 

systemic infections, and the affected birds may 

appear well-nourished and have minimal external and internal abnormalities. Salmonellosis in 

shorebirds may also present as subacute to chronic 

infections, and the birds are emaciated. Caseous 

granulomas in the esophagus and crop are not a 

feature of systemic salmonellosis of shorebirds. 

Proportion of diagnostic cases

0.0

0.2

0.4

0.6

0.8

1.0

Occurrence of salmonellosis in wild bird carcasses, by family.

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 13. Occurrence of salmonellosis in wild 

birds 2001–10, by order.
Selected Wildlife Diseases 13

EXPLANATION

Salmonellosis, as a percentage 

of total avian cases 

investigated at NWHC

0

1 to 15

16 to 33

34 to 67

68 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VA

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VA

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 14. Prevalence of salmonellosis diagnosed in wild bird disease outbreaks, as a percentage of 

all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
14 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

0

5

10

15

20

25

30

Number of diagnostic cases

Year

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 15. Annual number of cases of salmonellosis in wild birds.

0

5

10

15

20

25

30

Graph showing the monthly number of cases of salmonellosis in wild bird carcasses.

Month

Number of diagnostic cases

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Figure 16. Monthly number of cases of salmonellosis in wild 

birds.

identified S. typhimurium from wild birds, must be considered 

zoonoses that are potential pathogens to all amphibians, reptiles, birds, and mammals. Because persons may be infected 

by most serotypes of salmonellae, it is recommended that all 

persons who handle sick and dead birds around or near their 

birdfeeders wear disposable gloves or place a plastic bag on 

their hand when collecting carcasses. Furthermore, handlers of 

dead birds are urged to practice excellent personal hygiene by 

thoroughly washing their hands after handling dead birds and 

bird-feeding stations, and they should remove soiled clothing promptly and wash it. Persons with birdfeeders and dead 

birds in their yard also are urged to keep their pets away from 

sick and dead birds, and remove, wash, and disinfect feeding 

stations weekly. Removing or closing bird-feeding stations 

is recommended when morbidity and mortality persists after 

washing and disinfecting the feeder. 

Selected References

Friend, M., 1999, Salmonellosis, chap. 9 of Friend, M., and 

Franson, J.C., eds., Field manual of wildlife diseases: general field procedures and diseases of birds, U.S. Geological 

Survey Information and Technology Report 1999–001 

p. 99–109.

Daoust, P., and Prescott, J.F., 2007, Salmonellosis, chap. 13 

of Thomas, N.J., Hunter, D.B., and Atkinson, C.T., eds., 

Infectious diseases of wild birds: Ames, Iowa, Blackwell 

Publishing, p. 270–288.

Significance and Zoonotic Risk

All serotypes of salmonellae, including the commonly 
Selected Wildlife Diseases 15

Exotic Trematode Infections

Cause or Etiology

In the U.S., three introduced trematodes that were 

originally native to Europe cause lethal intestinal infections in 

endemic waterfowl and shorebirds: Cyathocotyle bushiensis, 

Leyogonimus polyoon, and Sphaeridiotrema globulus. These 

three exotic trematodes were first detected in 1960s along the 

St Lawrence River and lower Great Lakes (Gagnon, 1990); 

in 1996 in Wisconsin; and in 1928 along the Potomac River. 

Their introduced exotic intermediate host, also from Europe, 

is the aquatic snail, Bithynia tentaculata. Most native aquatic 

snails have proven unsuccessful as intermediate hosts for the 

three introduced trematodes. 

Host Species

Lethal intestinal infections by C. bushiensis, L. polyoon, and S. globulus are most commonly detected in lesser 

scaup ducks and American coots (fig. 17; Fulica americana) 

(Herrmann and Soreson, 2011). Other waterfowl that may be 

infected by C. bushiensis and S. globulus and that usually are 

found dead with scaups and coots are ruddy ducks (Oxyura 

jamaicensis), redhead ducks (Aythya ferina), mallards (Anas 

platyrhynchos), and blue-winged teal (Anas discors). C. 

bushiensis and S. globulus uncommonly are a cause of death 

in bufflehead ducks (Bucephala albeola), canvasback ducks 

(Aythya valisineria), ring-necked ducks (Aythya collaris), 

northern shoveler ducks (Anas clypeata), white-winged scoters 

(Melanitta fusca), Canada geese (Branta canadensis), snow 

geese (Chen caerulescens), and swans. 

The life cycle of the three parasites begins with excretion 

of parasite eggs by birds in their feces. The trematode eggs 

hatch and release the motile form of the parasite, the miracidia, into the aquatic environment. Free-swimming miracidia 

invade aquatic snails, usually Bithynia tentaculata. Life cycles 

of C. bushiensis, L. polyoon and S. globulus involve one or 

more intermediate hosts. The first intermediate host for these 

parasites is an introduced aquatic snail, Bithynia tentaculata, 

but S. globulus also may use other snail hosts including Elimia 

virginica, Fluminicola virens, and Oxytrema silicula (the latter two snails are native to North America) and pass through 

multiple stages of development, eventually producing cercariae. Cercariae of C. bushiensis and S. globulus are released 

by the snail into the water; the free-swimming cercariae then 

invade and reinfect the individual snail and nearby snails. The 

cercariae develop into metacercariae in the snail; the metacercariae are the infectious stage when eaten by birds. 

The life cycle for L. polyoon is more complicated, 

with second intermediate hosts that include aquatic insects 

and amphipods. When Bithynia snails release cercariae of 

L. polyoon, the free-swimming cercariae may invade numerous intermediate hosts including amphipods and a variety of 

aquatic insects. The cercariae develop into infectious metacercariae in these invertebrates. In addition, certain predatory 

aquatic insects, such as caddis fly larvae (order: Tricoptera) 

and damsel fly larvae (order: Odonata; suborder: Zygoptera) 

may accumulate live, infectious metacercariae into their 

bodies by consuming other infected invertebrates. 

American coots (F. americana) probably become 

infected by L. polyoon by eating caddis flies and 

damsel flies. Morbidity and mortality associated 

with these exotic trematodes is generally limited to 

wetlands where B. tentaculata is present.

Geographic Distribution

Presently, C. bushiensis, L. polyoon, and 

S. globulus have a limited distribution in North 

America (fig. 18). S. globulus has the largest 

introduced range: it is found along the Potomac 

River in Maryland; the Hudson River in New York; 

the St. Lawrence River in Canada and the United 

States (Gibson and others, 1972); the southern 

Great Lakes; at least three small inland lakes in 

Wisconsin (Cole, 2001); the Upper Mississippi 

River (Sauer and others, 2007), which borders 

the States of Iowa, Wisconsin and Minnesota; 

along the Mississippi River in eastern and central 

Minnesota; and in three or four lakes in western 

Montana. Infections by C. bushiensis occur in the 

same regions as S. globulus, but they have not 

been reported on the Potomac and Hudson Rivers. 

L. polyoon has a limited distribution in Wisconsin 

and Minnesota, but because the parasite is present 

Proportion of diagnostic cases

0.0

0.2

0.4

0.6

0.8

1.0

Exotic trematode infections in wild birds ....

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 17. Occurrence of exotic trematode infections in wild birds 2001–10, 

by order.
16 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

EXPLANATION

Trematodes, as a percentage 

of total avian cases 

investigated at NWHC

0

1 to 27

28 to 30

31 to 35

36 to 50

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Prevalence of exotic trematode infections diagnosed in wild bird disease outbreaks, as a percentage of all wild 

bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 18. Prevalence of exotic trematode infections diagnosed in wild bird disease outbreaks, as a 

percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

in the Upper Mississippi 

River, continued 

northerly and southerly 

spread along the river is 

anticipated. The spread 

of C. bushiensis and L. 

polyoon with infection 

of waterfowl and shore 

birds, however, must be 

preceded by the spread of 

the exotic host snail, B. 

tentaculata.

Seasonality

Although these 

parasites may be found in 

small mortality events in 

every month of the year 

(fig. 20), larger mortality 

events occur in midspring and mid-autumn 

and are each associated 

with northern and 

southern migrations of 

waterfowl, respectively. 

Many of the mortality 

events in spring and 

autumn occurred on the 

upper Mississippi River 

where waterfowl stop for 

a few days to rest and 

feed before resuming 

their migrations. 

Mortality rates of up to 

11,000 coots at one site 

in one season have been 

associated with these 

trematode parasites in 

Wisconsin (Cole, 2001). 

Signs and Lesions

Field and clinical 

signs of infection by 

exotic trematodes may be 

confused with botulism, 

because general weakness 

and flaccid paralysis are 

often observed. Some 

sick birds may show 

weak flight or dive 

attempts, a tendency to 

remain behind and swim 

into cover when the rest 

of flock would flush, and 

may quickly become 
Selected Wildlife Diseases 17

exhausted when trying to escape. Infrequently, a bird may pass 

bloody or black feces. Affected sick and dead birds usually 

are in fair to good body condition, consistent with reports 

of mortality occurring 3–8 days after infection. Internally, 

C. bushiensis has a tropism for the caeca, but the trematode 

occasionally is present in the distal ileum and colon. Apices of 

the caeca may show hemorrhage and diphtheritic or caseous 

ulcerations in the mucosa. L. polyoon has a tropism for the 

duodenum and jejunum, where it may cause hemorrhage into 

the gut lumen. S. globulus has a tropism for the jejunum and 

ileum, but in heavy infections of more than 3,000 trematodes, 

0

10

20

30

40

50

60

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 19. Annual number of cases of exotic trematode infection 

in wild birds.

Month

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

0

10

20

30

40

50

Number of diagnostic cases

Graph showing the monthly number of cases of exotic trematode 

Figure 20. Monthly number of cases of exotic trematode 

infection in wild birds.

the trematodes may also be present in the duodenum and 

caeca; hemorrhage into the lumen and mild flaccidity and 

dilation of the wall of the intestine may be present. Fewer than 

10 adult C. bushiensis have been associated with deaths of 

waterfowl. C. bushiensis and S. globulus may cause morbidity 

and mortality as single infections, but it is common for 

both to be present in the same bird. L. polyoon has a limited 

geographic distribution restricted to Minnesota and Wisconsin; 

hence, it usually is found in the presence of the other two 

species of trematodes.

Significance and Zoonotic Risk

There is no zoonotic risk to humans and pets by these 

exotic trematodes, but infections of domestic breeds of ducks 

and geese are possible. Infections by these trematodes are 

most common in waterfowl and shorebirds such as scaups and 

coots, which actively feed on aquatic invertebrates, and are 

less common in waterfowl such as geese and swans, which 

generally feed on vegetation. 

Selected References

Cole, R.A., 2001, Exotic parasite causes large scale mortality in American coots: accessed on May 30, 2012, at http://

www.nwhc.usgs.gov/publications/fact_sheets/pdfs/fact_

lpolyoon.pdf.

Cole, R.A., Roderick, C.L., Taft, S., Choudhury, A., Kanev, 

I., and Brockman-Mederas, K.M., 2000, Life cycle of 

Leyogonimus polyoon (Trematoda: Lecithodendriidae): 

presentation at Joint Meetings of the American Society of 

Parasitologists and Society of Protozoologists, San Juan, 

Puerto Rico. June 24–28, 2000.

Gagnon, C., 1990, Pathological consequences of infection by 

Cyathocotyle bushiensis Khan, 1962, and Sphaeridiotrema 

globulus (Rudolphi, 1814) in two species of dabbling ducks: 

Montreal, Quebec, Canada, McGill University, M.S. thesis, 

167 p.

Gibson, G.G., Broughton, E., and Choquette, L.P.E., 1972, 

Waterfowl mortality caused by Cyathocotyle bushiensis

Khan, 1962 (Trematoda: Cyathocotylidae), St. Lawrence 

River, Quebec: Canadian Journal of Zoology, v. 50, no. 11, 

p. 1351–1356. 

Herrmann, K.K., and Sorenson, R.E., 2011, Differences in 

natural infections of two mortality-related trematodes in 

lesser scaup and American coot: Journal of Parasitology v. 

97, no. 4, p. 555–558.

Sauer, J.S., Cole, R.A., and Nissen, J.M., 2007, Finding the 

exotic faucet snail (Bithynia tentaculata): investigation of 

waterbird die-offs in the Upper Mississippi River National 

Wildlife and Fish Refuge: U.S. Geological Survey OpenFile Report 2007–1063, 3 p. 
18 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

Lead Poisoning (Plumbism)

Cause or Etiology

Lead poisoning is a persistent disease that is declining 

due to a ban on the use of lead shot in hunting waterfowl in 

1991. Lead shot pellets are persistent in the environment, 

however, so ingestion of lead remains a problem in numerous 

areas where waterfowl were hunted throughout most of the 

20th century. Lead poisoning also is associated with accidental 

ingestion of lead fishing sinkers by fish-eating birds (Franson 

and others, 2003) and ingestion by eagles and other avian 

scavengers of lead bullet fragments in the discarded entrails 

of hunter-killed deer and other big game animals. Infrequent 

cases of lead poisoning are associated with ingestion of chips 

and flakes of lead-based paint from old buildings and ingestion 

of mine wastes or other lead-based objects. In the U.S., some 

States have implemented local or regional bans on lead bullets 

and lead fishing tackle, but legislation to ban lead bullets and 

fishing tackle nationwide has not been passed. The use of lead 

shot is inconsistently banned for hunting upland gamebirds, 

including doves, so lead poisoning following ingestion of lead 

shot in terrestrial birds remains a problem (Franson and others, 

2009). 

Host Species and Annual Distribution

Lead poisoning affects a wide range of bird species, 

but in the years of this report, more than 95 percent of cases 

involved waterfowl (ducks, geese, and swans) and eagles 

(95 percent bald eagles; 5 percent golden eagles) (fig. 21). 

Other lead poisoned birds included three common loons, two 

sandhill cranes (Grus canadensis), a cowbird, and a black 

vulture (Coragyps atratus). The number of lead poisoning 

cases declined by 38 percent between 2001–5 (170 cases) 

and 2006–10 (96 cases). The data may be skewed towards 

submission of eagles, (81 percent of all cases between 2001 

and 2005 and 69 percent of all cases between 2006 and 2010 

were eagles).

Geographic Distribution

Lead poisoning is a nationwide problem (fig. 22), and 

only seven States (Hawai’i, Nevada, Alabama, Georgia, 

Delaware, Massachusetts, and Rhode Island) had no confirmed 

cases in 2001–10; however, lead poisoning had been detected 

in birds in several of the seven States in the prior years. The 

number of birds submitted from some of the seven States also 

was limited. 

Seasonality 

Cases of lead poisoning usually 

peak during the months of late autumn, 

winter, and early spring (November–

May) (fig. 24). Cases of lead poisoning 

occur year-round, but they are decidedly 

fewer during late spring, summer, and 

early autumn. Cases in late autumn and 

early winter are associated with duck and 

deer hunting seasons, and cases in midwinter and spring probably are due to 

birds ingesting lead shot on and near their 

wintering sites. 

Proportion of diagnostic cases

0.0

0.2

0.4

0.6

0.8

1.0

 Occurrence of lead poisoning in wild birds ...

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 21. Occurrence of lead poisoning in wild birds 2001–10, by order.
Selected Wildlife Diseases 19

EXPLANATION

Lead poisoning, as a percentage 

of total avian cases 

investigated at NWHC

0

1 to 30

31 to 50

51 to 75

76 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 19. Maps showing the prevalence of lead poisoning diagnosed in wild birds, as a percentage of all wild 

bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 22. Prevalence of lead poisoning diagnosed in wild birds, as a percentage of all wild bird 

disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
20 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

0

10

20

30

40

50

60

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 23. Annual number of cases of lead poisoning in wild 

birds.

Month

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

0

10

20

30

40

50

Number of diagnostic cases

Monthly number of cases of lead poisoning in wild birds. Figure 24. Monthly number of cases of lead poisoning in wild 

birds.

Signs and Lesions

Most birds with lead poisoning have a chronic 

intoxication and are moderately to severely emaciated. 

Uncommon cases of lead poisoning in well-nourished birds 

are considered to be examples of acute lead poisoning. In 

the field, poisoned birds are weak, ataxic, fearless, and may 

be reluctant to fly or fly weakly. Affected Canada geese 

may have abnormal crooked postures of the head and neck 

during flight, and a marked change in tone of the call is 

occasionally evident. Waterfowl also may show abnormal 

postures or paralysis of the wings. At necropsy, lead poisoned 

birds usually have no body fat and severe atrophy of breast 

(pectoral) muscles. Distention or impaction of the esophagus 

and proventriculus with feed is noted in less than 25 percent of 

poisoned waterfowl. Offending lead objects may be detected 

in the lumen of the gizzard. 

Significance and Zoonotic Risk

Risk of secondary lead poisoning in predators, 

scavengers, pets, and humans is highly variable. The amount 

of lead in tissues, for example, the muscles and liver, of lead 

poisoned waterfowl is considered insufficient for a person 

to develop clinical signs of poisoning by eating one or a few 

birds. Consumption of a large number of poisoned birds in a 

few days would be necessary for toxic levels to accumulate 

in humans; however, consumption of lead shot in the flesh of 

waterfowl may lead to clinical lead poisoning in children and 

adults. Predators and scavengers may become secondarily 

poisoned by ingesting lead shot and bullet fragments in the 

gizzards, flesh, or entrails of animals. 

Selected References

Franson, J.C., Hansen, S.P., Creekmore, T.E., Brand, C.J., 

Evers, D.C., Duerr, A.E., and DeStefano, S., 2003, 

Lead fishing weights and other fishing tackle in selected 

waterbirds: Waterbirds, v. 26, p. 345–352. 

Franson, J.C., Hansen, S.P.,; and Schulz, J.H., 2009, Ingested 

shot and tissues lead concentrations in mourning doves, in, 

Watson, R.T.; Fuller, M.; Pokras, M.; and Hunt, W.G., eds., 

Ingestion of lead from spent ammunition: implications for 

wildlife and humans: Boise, Idaho, The Peregrine Fund, 

p. 1–12. 

Pain, D.J., Cromie, R.L., Newth, J., Brown, M.J., Crutcher, 

E., Hardman, P., Hurst, L., Mateo, R., Meharg, A.A., 

Moran, A.C., Raab, A., Taggart, M.A., and Green, R.E., 

2010, Potential hazard to human health from exposure 

to fragments of lead bullets and shot in the tissues of 

game animals: PLOS ONE, v. 5, no. 4, p. 1–17, accessed 

November 29, 2012, at http://www.plosone.org/article/

info:doi%2F10.1371%2Fjournal.pone.0010315

The Wildlife Society and American Fisheries Society, 2008, 

Sources and implication of lead ammunition and fishing 

tackle on natural resources: Technical Review 08–01, 62 p.
Selected Wildlife Diseases 21

Virulent Newcastle Disease Virus

Cause or Etiology

Virulent Newcastle disease virus (vNDV) belongs in 

the genus, Rubulavirus, in the viral family, Paramyxoviridae, 

and Subfamily, Paramyxovirinae. Several strains of NDV 

have been identified, but only some strains are designated 

“virulent.” Isolates of vNDV are considered virulent only 

when poultry (chickens) are affected and may or may 

not be virulent to wild birds. Virulent and nonvirulent 

isolates of NDV are considered one species of virus: avian 

paramyxovirus-1 (APMV–1). This virus has a ribonucleic acid 

(RNA) genome and other viruses in the genus, Rubulavirus, 

include human mumps virus, avian paramyxovirus-2 through 

9, human parainfluenza virus-2 and 4, three simian and porcine 

viruses, and probably Hendra virus of fruit bats, horses, and 

humans in Australia. Isolates of vNDV and NDV in recent 

years have been named cormorant paramyxovirus, pigeon 

paramyxovirus, and others. Each isolate of NDV, regardless of 

host species of bird, only is designated virulent (vNDV) when 

its virulence in chickens is established or a specific virulence 

gene is detected molecularly. 

Host Species and Annual Distribution

Between 2001and 2010, vNDV has been isolated almost 

exclusively from double-crested cormorants (Phalacrocorax 

auritus) in the U.S. (fig. 25). The vast majority of these 

isolates come from die-offs of nestling and fledgling 

cormorants from colonial nesting sites. Usually, these colonial 

nesting sites also have nesting pelicans, gulls, and other 

shorebirds, but vNDV has not been isolated from other species 

of birds at cormorant nesting sites. 

Geographic Distribution

Virulent Newcastle disease virus has been detected in 

die-offs of immature cormorants in seven States including 

South Dakota, Minnesota, Wisconsin, California, Maryland, 

Nevada, and North Dakota; however, over half of the cases 

of vNDV in cormorants have occurred in States bordering the 

Great Lakes (Wisconsin and Minnesota; fig. 26). Colonies 

of nesting cormorants also are common in North and South 

Dakota, where vNDV has been detected four times since 2001.

Isolates of APMV–1 from cormorants prior to 2001 were 

poorly characterized, so it is unknown when vNDV may have 

emerged as a disease in cormorants. 

Proportion of diagnostic cases

Occurrence of virulent Newcastle disease virus in wild bird carcasses, by family

0.0

0.2

0.4

0.6

0.8

1.0

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes 

Figure 25. Occurrence of virulent Newcastle disease virus in wild birds 2001–10, 

by order.
22 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

EXPLANATION

Virulent Newcastle disease 

virus, as a percentage of 

total avian cases 

investigated at NWHC

0

1 to 27

28 to 30

31 to 35

36 to 50

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 22. Maps showing the prevalence of virulent Newcastle disease virus diagnosed in wild bird disease 

outbreaks, as a percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 26 Prevalence of virulent Newcastle disease virus diagnosed in wild bird disease outbreaks, as 

a percentage of all wild bird disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Seasonality 

Because vNDV 

occurs predominantly 

in nestling and fledgling 

double-crested cormorants, 

there is a strong seasonal 

occurrence of the disease 

in months of the late spring 

and early summer. There is 

some variability in months 

of first occurrence that generally is related to weather 

conditions. About 75 

percent of cases of vNDV 

are reported in June, July, 

August, September, and 

October (fig. 28). 

Signs and Lesions

Nestling cormorants 

usually are found dead in 

or near their nests with no 

record of clinical signs. 

Fledgling cormorants also 

may be found dead, but 

also are detected because 

of neurological signs 

including severe lethargy, 

ataxia, or paralysis of one 

or more limbs. Infected 

nestlings may be emaciated or in fair to good body 

condition, while fledgling 

cormorants with neurological signs are more likely to 

be emaciated. No specific 

abnormalities are detected 

grossly in visceral organs 

of vNDV-infected cormorants, but histologically, 

many fledgling cormorants have minimal to 

moderate nonsuppurative 

encephalitis. 
Selected Wildlife Diseases 23

0

10

20

30

40

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

disease virus in wild birds.

Figure 27. Annual number of cases of virulent Newcastle 

0

10

20

30

40

50

Graph showing the monthly number of cases of virulent Newcastle 

disease virus in wild birds. 

Month

Number of diagnostic cases

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

disease virus in wild birds.

Figure 28. Monthly number of cases of virulent Newcastle 

Significance and Zoonotic Risk

Although there are rare reports of humans developing 

mild infections by NDV, human infections by vNDV from 

cormorants have not been documented. Although vNDV has 

only been detected in cormorants in the U.S. in the last 10 

years (2001–10), it is likely other wild birds are susceptible to 

infection and we remain alert for this occurrence. 

Selected References

Cattoli, G., Susta, L., Terregino, C., and Brown, C.; 2111, 

Newcastle disease: a review of field recognition and current 

methods of laboratory detection: Journal of Veterinary 

Diagnostic Investigation, v. 23, no. 4, p. 637–656.

Diel, D.G., Miller, P.J., Wolf, P.C., Mickley, R.M., Musante, 

A.R., Emanueli, D.C., Shively, K.J., Pedersen, K., and 

Afonso, C.L., 2012, Characterization of Newcastle disease 

viruses isolated from cormorant and gull species in the 

United States in 2010: Avian Diseases, v. 56, p. 128–133.

Hines, N.L., Killian, M.L., Pedersen, J.C., Reising, M.M., 

Mosos, N.A., Mathieu-Benson, C., and Miller, C.L., 2012, 

An rRT-PCR assay to detect the matrix gene of a broad 

range of avian paramyxovirus serotype-1 strains: Avian 

Diseases, v. 56, p. 387–395.
24 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

West Nile Virus

Cause or Etiology

West Nile virus (WNV) is in the genus, Flavivirus, in 

the family, Flaviviridae, which are all mosquito-borne viruses 

with RNA as their genome. Other viruses in the genus, 

Flavivirus, include Dengue virus, Japanese encephalitis virus, 

yellow fever virus, and several other tick-borne and mosquitoborne species of virus. The viral genomes that were examined 

from infected birds and humans in the U.S. suggest the strain 

of WNV introduced into the New York City area in 1999 may 

have originated from Israel (Lanciotti and others, 1999).

Host Species and Annual Distribution

West Nile virus has been isolated in cultures from 

many species of dead birds and a few squirrels. To date, 

WNV has not been isolated from any species of amphibians, 

nor any turtles and tortoises (chelonians). Mammals larger 

than squirrels are infrequently submitted to the NWHC, so 

we have not isolated WNV from native wild cats (felids), 

foxes and wolves (canids), raccoons (procyonids), and 

groundhogs, beavers, pikas, rabbits, and field mice (rodents 

and lagomorphs). The bird most commonly infected by WNV 

was the American white pelican (fig. 29; 75 of 172 cases, 

or 44 percent). Infected pelicans were mostly nestlings and 

fledglings in colonial nesting sites from northern States; adults 

were infrequently infected. Other species of birds at the same 

nesting sites, including gulls, terns, and cormorants, also Proportion of diagnostic cases

0.0

0.2

0.4

0.6

0.8

1.0

 Occurrence of West Nile virus in wild birds ...

Taxonomic order

Anseriformes

Charadriiformes

Ciconiiformes

Columbiformes

Falconiformes

Gaviiformes

Galliformes

Gruiformes

Passeriformes

Pelecaniformes

Podicipediformes

Strigiformes

Figure 29. Occurrence of West Nile virus in wild birds 2001–10, by order.

were infected but at much lower frequency. Raptors including 

owls, vultures, falcons, hawks, and eagles were the second 

most commonly infected group of birds (34 of 172, or 19.8 

percent); most infected raptors were adult or subadult birds. 

Most raptors also were emaciated, while pelicans and other 

shorebirds such as terns and gulls usually were in fair to good 

body condition, indicating the latter group of birds probably 

had acute systemic infections similar to those in crows, jays, 

and other corvids. It must be noted that these data include 

only cases submitted as part of mortality investigations; cases 

submitted as part of the WNV surveillance system, which 

largely consisted of corvids, are not included in this summary.

The specific mosquitoes involved in the transmission of 

WNV to wild birds were not studied when sick and dead birds 

were collected. 

Geographic Distribution

West Nile virus was first detected in dead American 

crows and zoological birds in New York City in 1999. Within 

months, the virus spread northwards and southwards to 

adjacent States. Within 5 years, the virus had spread westward 

to most of the contiguous 48 States (fig. 30). The spread of 

WNV into new States and counties often was marked by 

deaths of urban and suburban American crows (Caffrey and 

others, 2005; Eidson and others, 2001), although, WNV has 

been isolated from many species of passerines, pelicans, 

raptors, waterfowl, shorebirds, and others. WNV is now 

considered endemic in the lower 48 States but has not been 

detected in Alaska or Hawaii.

Seasonality

Occurrence of infection by WNV 

is strongly linked to time of the year 

(fig. 32). Morbidity and mortality 

associated with WNV generally occurs 

annually a few weeks following 

emergence of large populations of 

mosquitoes. The first fatal cases of 

infection by WNV generally occur in 

July or August, but cases may appear 

months earlier in southern States. 

Generally, cases of WNV decline 

abruptly and sharply following the 

first freezing weather in the autumn. 

Detection of infections by WNV in 

winter months is rare. 
Selected Wildlife Diseases 25

EXPLANATION

West Nile Virus, as a 

percentage of total avian 

cases investigated at NWHC

0

1 to 6

7 to 17

18 to 25

26 to 35

36 to 67

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Maps showing the prevalence of West Nile virus in wild bird disease outbreaks, as a percentage of all wild bird 

disease outbreaks, by State, for A, 2001–5 and B, 2006–10. 

Figure 30. Prevalence of West Nile virus in wild bird disease outbreaks, as a percentage of all wild bird 

disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Signs and Lesions

Passerine birds, 

including corvids, 

generally are found dead 

and signs prior to death 

are seldom observed. 

Immature pelicans, gulls, 

terns, and cormorants 

with infections by WNV 

are usually found dead, 

but a small number 

are found collapsed, 

semicomatose, or very 

lethargic. Raptors with 

infection by WNV 

usually are adults or 

subadults. Neurological 

signs predominate 

in infected raptors, 

including head tilt, 

limb paralysis, ataxia, 

and inability to fly or 

walk. Most raptors 

also are emaciated and 

have nonsuppurative 

encephalitis, while most 

passerines are in fair to 

good body condition. 

Significance and 

Zoonotic risk

Direct transmission 

of WNV from birdto-bird or bird-tomammal is rare and 

generally involves 

laboratory accidents, 

failure of laboratory 

staff to use personal 

protective equipment, 

or infrequently, bird or 

mammal scavenging or 

consumption of WNVinfected carcasses. 
26 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

0

10

20

30

40

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 31. Annual number of cases of West Nile virus in wild 

birds.

0

10

20

30

40

50

Monthly number of cases of West Nile virus in wild birds.

Month

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Number of diagnostic cases

Figure 32. Monthly number of cases of West Nile virus in wild 

birds.

Although WNV is considered a zoonotic disease, the major 

hazard to persons probably is from exposure to infected 

mosquitoes while doing field work. Although pet dogs 

and cats can be exposed to WNV if they consume infected 

carcasses, most cases of WNV infection in pets probably are 

owing to transmission by bites of mosquitoes. Many States 

and counties begin soliciting the collection and submission 

of dead passerine birds from urban and suburban locations in 

the spring and then publicize the first detection of WNV in a 

dead bird each year to alert citizens of the need to apply insect 

repellants to avoid infection while outdoors. 

Selected References

Caffrey, C., Smith, S.R.C., and Weston, T.J., 2005, West Nile 

virus devastates an American crow population: The Condor, 

v. 107, p. 128–132. 

Dusek, R.J.; McLean, R.G.; Kramer, L.D.; Ubico, S.R.; 

Dupuis, A.P., II; Ebel, G.D.; and Guptill, S.C., 2009, 

Prevalence of West Nile virus in migratory birds during 

spring and fall migration: American Journal of Tropical 

Medicine and Hygiene, v. 81, no. 6, p. 1151–1158.

Eidson, M., Kramer, L., Stone, W., Hagiwara, Y., Schmitt, K., 

and the New York State West Nile Virus Avian Surveillance 

Team, 2001, Dead bird surveillance as an early warning 

system for West Nile virus: Emerging Infectious Diseases, 

v. 7, p. 615–620. 

Hofmeister, E.K., 2011, West Nile virus: North American 

experience: Integrative Zoology, v. 6, p. 279–289.

Lanciotti, R.S., Roehrig, J.T., Deubel, V., Smith, J., Parker, 

M., Steele, K., Volpe, K.E., Crabtree, M.B., Scherret, 

J., Hall, R., Mackenzie, J., Cropp, C.B., Panigraphy, B., 

Malkinson, M., Komar, N., Savage, H.M., Stone, W., 

McNamara, T., and Gubler, D.J., 1999, Origin of the West 

Nile virus responsible for an outbreak of encephalitis in the 

northeastern United States: Science, v. 286, p. 2333–2337. 

McLean, R.G., and Ubico, S.R., 2007, Arboviruses in birds, 

chap. 2 of Thomas, N.J.; Hunter, D.B.; and Atkinson, 

C.T., eds., Infectious diseases of wild birds: Ames, Iowa, 

Blackwell Publishing, p. 17–62. 

Weingartl, H.M., Neufeld, J.L., Copps, J., and Marszal, P., 

2004, Experimental West Nile virus infection in blue jays 

(Cyanocitta cristata) and crows (Corvus brachyrhynchos): 

Veterinary Pathology, v. 41, p. 362–370. 

Wunschmann, A., Shivers, J., Carroll, L., and Bender, J., 

2004, Pathological and immunohistochemical findings 

in Americans crows (Corvus brachyrhynchos) naturally 

infected with West Nile virus: Journal of Veterinary 

Diagnostic Investigation, v. 16, p. 329–333. 
Selected Wildlife Diseases 27

Selected Herpetological Diseases

Amphibian Chytridiomycosis

Cause or Etiology

The pathogenic chytrid fungus of amphibians is 

Batrachochytrium dendrobatidis (“Bd”). There probably 

is no other single disease agent in human records that has 

caused more global deaths, decimations, extirpations, and 

extinctions than Bd has caused to the order Anura, which 

includes frogs and toads (Skerratt and others, 2007). Chytrid 

fungi are a distinct phylum of protists called Chytridiomycota, 

and Bd is the only species of chytrid fungus to parasitize or 

infect vertebrate animals. Numerous taxa of chytrid fungi 

are ubiquitous in the environment, especially wetlands. 

Although other genera of chytrid fungi break down keratin 

proteins found in skin, hair, feathers, and claws or insect 

exoskeletons (chitin) in the environment, Bd of amphibians 

is an infection strictly of keratinized epidermal cells, and thus 

chytridiomycosis has not been detected in fish, reptiles, birds, 

or mammals. Chytrid fungi cause intracellular infections 

and form spherical to ovoid thalli (microscopic single-cell 

structures); chytrid fungi do not form hyphae like higher 

fungi. The infectious stage of Bd is a uniflagellate zoospore 

that forms in, and is released from, the thalli. Resting stages or 

environmentally hardy spores have not been described, but it 

is not clear how the organism seems to persist in wetlands year 

after year without suitable amphibian hosts or through drying 

of wetlands. Proportion of disease events

0.0

0.2

0.4

0.6

0.8

1.0

Frog Salamander 

and newt

Toad Treefrog Turtle

Occurrence of chytridiomycosis in amphibian s ...

Family

Figure 33. Occurrence of chytridiomycosis in amphibians, by family.

Host Species

Chytridiomycosis is an infection of amphibian larvae 

and postmetamorphs, but it is lethal only in postmetamorphic 

frogs and toads (fig. 33). During metamorphosis, the epidermis 

of amphibians changes from keratin-free larval skin to 

keratinized adult skin. The only keratin in larval frogs and 

toads occurs in the oral disc; the black jaw sheaths and tooth 

rows are the only keratin in tadpoles. Larval salamanders 

may have traces of keratin at the tips of their digits, but 

lethal infections of larval amphibians are not reported, 

probably because of their very limited amount of keratin. It 

is likely that where chytridiomycosis occurs in the larvae, 

very few amphibians can complete metamorphosis quickly 

enough to escape chytrid infection of their newly forming 

keratinized adult skin. Bd infections are rarely lethal in adult 

salamanders with the exception of eastern red-spotted newts 

(Notophthalmus viridescens) and most genera of salamanders 

in Central and South America. Susceptibility of caecilians 

(legless amphibians) is largely unknown. 

Geographic Distribution

Batrachochytrium dendrobatidis is found on all six 

continents that have endemic amphibians and also on many 

large and small island nations. Based on histological studies of 

museum specimens in the northern and western United States 

and western Canada, this infection first appeared in the late 

1960s and early 1970s. It was the probable cause of massive 

amphibian population declines in the 1970s and then extirpation of populations and extinctions of 

many species in subsequent decades. Bd 

was not identified, however, until the mid 

to late 1990s (Berger and others, 1998), 

which may have allowed its international 

dissemination. The geographic origin of 

Bd is unknown, although there is some 

evidence that the pathogen is endemic 

to southern Africa (Weldon and others, 

2004) and may have been spread globally by the importation of millions of live 

wild-borne African clawed frogs (Xenopus 

laevis) for research and clinical studies. 

In semitropical, tropical, and neotropical locations, chytridiomycosis tends 

to be distributed by elevation. Amphibian 

species and populations above 200–400 

meters tend to be much more susceptible 

to chytridiomycosis than species found 

at lower elevations, probably because of 

cooler temperatures at higher elevations. 

Because Bd is psychrophilic and ceases to 

grow at temperatures above 30–32 °C, the 

infection may be inhibited at hotter, lower 

elevations in tropical regions (Woodhams 

and others, 2003); however, Bd also is 

killed by freezing. 
28 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

EXPLANATION

Chytridiomycosis, as a 

percentage of total amphibian 

cases investigated at NWHC

0

1 to 25

26 to 50

51 to 75

76 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 28. Maps showing the prevalence of chytridiomycosis diagnosed in amphibians, as a percentage of all 

amphibian disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Figure 34. Prevalence of chytridiomycosis diagnosed in amphibians, as a percentage of all amphibian 

disease outbreaks, by State, for A, 2001–5 and B, 2006–10.

Seasonality 

Chytridiomycoses 

(lethal and nonlethal 

infections) have been 

detected in amphibians 

in every month (fig. 36), 

but lethal infections of 

postmetamorphs tend to 

occur during cool weather 

rather than cold or hot 

weather. Hence, in southwestern States, where 

summers usually are hot 

(greater than 30 °C), sick 

and dead amphibians 

usually are detected in the 

late autumn, winter, and 

early spring. In temperate 

regions, the infection may 

be seen throughout the 

late spring, summer, and 

autumn. At higher elevations, where snowmelt 

may not be complete until 

late spring, chytridiomycosis is mostly detected 

during summer months. 

In temperate zones and 

high altitudes, mortality events may occur 

promptly following the 

completion of metamorphosis in May, June, 

and July. Some tadpoles 

may persist as larvae for 

one or two winters, and 

infections in tadpoles may 

be detected year-round—

even in those collected 

beneath ice-covered lakes. 

Signs and Lesions

Mortality events 

due to chytridiomycosis 

usually are insidious and 

often pass undetected. 

The paucity of observed 

casualties is greatly 

disproportionate to the 

severity of the population 

decline. In tadpoles (the 

anuran larval stage), the 

infection is limited to specific structures of the oral 

disc and palate. Normally, 
Selected Wildlife Diseases 29

the jaw sheaths and tooth rows of tadpoles are black; with chytrid infection, there is a loss of the black keratinized cells, and 

these structures show varying degrees of white discoloration. 

Mortalities associated with oral chytridiomycosis in tadpoles 

do not occur. In postmetamorphic frogs and toads, chytrid 

infections may begin promptly in newly formed keratinized 

adult skin. Mortality rates in recently metamorphosed frogs 

and toads may be exceedingly high. Infected postmetamorphs 

are lethargic, fearless, and may have a poor righting reflex. 

The infection is most intense in the ventral skin, digits, and 

toe webs, and is absent or minimal in the dorsal skin. Some 

infected frogs and toads may show abnormal molt (dysecdysis) in which flakes of unshed skin remain attached to the 

body or the tips of digits. In advanced infections, the ventral 

skin may appear mildly thickened and light grayish-brown. 

Because amphibians do not drink water but absorb it through 

their skin, it is suspected that accumulation of numerous layers 

2

4

6

8

10

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

0

Figure 35. Monthly number of cases of chytridiomycosis in 

amphibians.

Figure 36. Monthly number of cases of chytridiomycosis in 

amphibians.

of unshed skin may inhibit water absorption; consequently, 

amphibians with severe infections often are found along shorelines or in surface water. Cause of death in chytridiomycosis 

is attributed to disruption of cutaneous functions resulting 

in lethal fluctuations in plasma electrolytes and subsequent 

cardiac arrest (Voyles and others, 2010).

Significance and Zoonotic Risk

Infection by Bd is a global epizootic, but the infection 

is limited to amphibians. There are no reports of Bd infection 

in any other class of vertebrates such as fish, reptiles, endotherms, or humans. 

Selected References

Berger, L., Speare, R., Daszak, P., Green, D.E., Cunningham, 

A.A., Goggin, C.L., Slocombe, R., Ragan, M.A., Hyatt, 

A.D., McDonald, K.R., Hines, H.B., Lips, K.R., Marantelli, 

G., and Parkes, H., 1998, Chytridiomycosis causes amphibian mortality associated with population declines in the 

rainforests of Australia and Central America: Proceedings 

of the National Academy of Sciences of the United States of 

America, v. 95, no. 15, p. 9031–9036. 

Berger, L., Speare, R., Hines, H.B., Marantelli, G., Hyatt, 

A.D., McDonald, K.R., Skerratt, L.F., Olsen, V., Clarke, 

J.M., Gillespie, G., Mahony, M., Sheppard, N., Williams, 

C., and Tyler, M.J., 2004, Effect of season and temperature 

on mortality in amphibians due to chytridiomycosis: Australian Veterinary Journal, v. 82, no. 31–36.

Hyatt, A.D., Boyle, D.G., Olsen, V., Boyle, D.B., Berger, L., 

Obendorf, D., Dalton, A., Kriger, K., Hero, M., Hines, H., 

Phillott, R., Campbell, R., Marantelli, G., Gleason, F., and 

Colling, A., 2007, Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis: 

Diseases of Aquatic Organisms, v. 73, p. 175–192. 

Skerratt, L.F., Berger, L., Speare, R., Cashins, S., McDonald, 

K.R., Phillott, A.D., Hines, H.B., and Kenyon, N., 2007, 

Spread of chytrdiomycosia has cause the rapid global 

decline and extinction of frogs: EcoHealth, v. 4, p. 125–134.

Voyles, J., Young, S., Berger, L., Campbell, C., Voyles, W.F., 

Dinudom, A., Cook, D., Webb, R., Alford, R.A., Skerratt, 

L.F., and Speare, R., 2010, Pathogenesis of chytridiomycosis, a cause of catastrophic amphibian declines: Science, v. 

326, p. 582–585.

Weldon, C., du Preez, L.H., Hyatt, A.D., Muller, R., and 

Speare, R., 2004, Origin of the amphibian chytrid fungus: 

Emerging Infectious Diseases, v. 10, p. 2100–2105.

Woodhams, D.C., Alford, R.A., and Marantelli, G., 2003, 

Emerging disease of amphibians cured by elevated body 

temperature: Diseases of Aquatic Organisms, v. 55, p. 

65–67. 

0

2

4

6

8

10

Month

Number of diagnostic cases

Monthly number of cases of chytridiomycosis in 

amphibian carcasses.

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.
30 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

Ranavirus

Cause or Etiology

Ranaviruses are deoxyribonucleic acid (DNA)-based 

viruses of the genus, Ranavirus, in the family, Iridoviridae. 

Occasionally, amphibian ranaviruses are called iridoviruses, 

but because there is a genus of insect viruses called Iridovirus, 

this practice is to be avoided. The type species of amphibian 

ranavirus is frog virus-3 (FV–3) which was first isolated 

and identified in the 1960s. Multiple strains of amphibian 

ranaviruses have been reported; most noteworthy are FV–3 

and Ambystoma tigrinum (tiger salamander) ranavirus. Other 

strains or isolates of ranaviruses often are named, for example, 

Bohle virus in Australia, Redwood Creek virus in northern 

California, and Southern High Plains virus, but the status of 

most isolates as species of Ranavirus has yet to be clarified. 

Ranaviruses also cause morbidity and mortality in fish and 

reptiles. Other viruses in the family, Iridoviridae, mostly infect 

insects and fish. 

Host Species

Mortality events due to ranaviruses occur most 

commonly in aquatic larval amphibians such as mole 

salamanders (Ambystoma spp.), true frogs (Rana spp.), and 

chorus frogs (Pseudacris spp.) (fig. 37). Infrequent isolates 

have been obtained from adult newts (Notophthalmus 

viridescens), adult treefrogs (Hyla spp.), and postmetamorphic Columbia spotted frogs (Lithobates [formerly 

Rana] luteiventris). At this laboratory, ranaviruses have been 

isolated from 16 species of frogs, 1 species of toad, and 6 

species of salamanders. In States east of the Mississippi River, 

especially the Atlantic coastal States, mortality events tend to 

involve all species within the affected wetland, for example, 

frogs, toads, and salamanders; in the Western States, where 

amphibian species are less diverse, mortality events tend to 

involve only one species per event. 

Geographic Distribution

Amphibian ranaviruses probably are present in every 

State with the possible exception of Hawai’i. In the 10 years 

of this report, mortality events attributed to ranaviruses were 

diagnosed in 25 States (fig. 38). Ranaviruses were detected 

in amphibians in some States, for example, Oklahoma, 

Louisiana, Georgia, New Jersey and Connecticut, where 

ranaviruses often were found in association with other 

infectious diseases. 

Seasonality 

Because mortality events due to ranaviruses mostly 

affect larval amphibians, there is a strong correlation 

between ranaviral die-offs and the presence of amphibian 

larvae in the spring and summer. Mortality events often 

are first detected in wetlands in April (fig. 40) when large 

populations of wood frogs (Lithobates sylvaticus, formerly 

Rana sylvatica) are present in eastern and northern wetlands. 

Early- and midsummer die-offs due to ranaviruses tend to 

involve a variety of amphibian species nationwide. Late 

summer and autumn mortality events involve mostly larval 

American bullfrogs (Lithobates catesbeianus, formerly Rana 

catesbeiana) and tiger salamanders.

Proportion of disease events

0.0

0.2

0.4

0.6

0.8

1.0

Occurrence of ranavirus in amphibian carcasses, ...

Frog Salamander 

and newt

Toad Treefrog Turtle

Family

Figure 37. Occurrence of ranavirus in 

amphibians and turtles, by family.
Selected Wildlife Diseases 31

EXPLANATION

Ranaviruses, as a percentage 

of total amphibian cases 

investigated at NWHC

0

1 to 20

21 to 40

41 to 50

51 to 100

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

TEXAS

NEW 

MEXICO

COLORADO

WYOMING

UTAH

NEVADA

CALIFORNIA

OREGON

WASH

IDAHO

MONTANA

NORTH 

DAKOTA

SOUTH 

DAKOTA

M I C H I G A N

OHIO

NEW YORK

FLORIDA

KY

CONN

NEW JERSEY

MASS

R I

MAINE VT

NH

DEL

VA MD

ARIZONA

ALASKA

HAWAII

IOWA NEBRASKA

OKLA

KANSAS

CANADA

MEXICO

MINN

WIS

ALA GA

WVA

SC

NC

LA

ARK

MO

ILL IND

PENN

MISS

TENN

A. 2001–5 

B. 2006–10 

Figure 31. Maps showing the prevalence of ranaviruses diagnosed in amphibian disease 

outbreaks, as a percentage of all amphibian disease outbreaks, by State, for A, 2001–5 and 

B, 2006–10.

Figure 38. Prevalence of ranaviruses diagnosed in amphibian disease outbreaks, as a percentage of all 

amphibian disease outbreaks, by State, for A, 2001–5 and B, 2006–10.
32 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

2

0

4

6

8

10

Year

Number of diagnostic cases

2001 2002 2003 2004 2005 2006 2007 2008 2009 2010

Figure 39. nnual number of cases of ranavirus in amphibians and 

turtles.

0

2

4

6

8

10

Month

Number of diagnostic cases

Jan. Feb. Mar. Apr. May June July Aug. Sept. Oct. Nov. Dec.

Figure 40. Monthly number of cases of ranavirus in amphibians 

and turtles.

Signs and Lesions

Field signs of a ranaviral epizootic are a sudden or explosive onset of morbidity and mortality in a wetland, often with 

hundreds or thousands of sick and dead amphibians in 1–5 

days. Mortality rates in a wetland usually exceed 90 percent. 

Affected individuals usually present with subtle-to-severe 

hemorrhages in the ventral skin, especially at the base of the 

hind limbs and around the vent opening. Hemorrhages may 

be present from tip of chin to tip of tail ventrally and may be 

petechial, paintbrush, or focally extensive. These hemorrhages 

have a tendency to fade with prolonged postmortem intervals 

or in frozen carcasses. Hence, descriptions of freshly dead 

amphibians in the field often are very helpful when examining frozen amphibians. Other clinical signs include lethargy, 

swimming erratically, weakly or on their sides, and mild-tosevere fluid accumulation under the skin in the lymphatic sacs 

of the abdomen and proximal hind limbs. Internally, there may 

be clear, fibrinous, or sanguineous fluid accumulation in the 

body cavity (called hydrocoelom), and petechial or paintbrush 

hemorrhages on the serosal surfaces of viscera, especially the 

heart, stomach, and liver. Occasionally, white minute foci of 

necrosis are evident in the liver or spleen. Ulcers of the skin 

and palate tend to be randomly scattered, but they are detected 

in a low percentage of casualties. 

Significance and Zoonotic Risk

At present, ranavirus infections appear to be limited 

to fish, amphibians, and reptiles, (ectothermic vertebrates). 

The virus generally cannot be cultured at temperatures above 

30 °C, so it probably is not transmissible to domestic animals 

and humans; however, ranaviruses will grow in bird and mammal cell lines that are incubated at temperatures less than 30 

°C. At present, ranaviral infections involving more than one 

class of vertebrates—either fish and amphibians or amphibians 

and reptiles—at a site are rarely reported in the wild. 

Selected References

Docherty, D.E., Meteyer, C.U., Wang, J., Mao, J., Case, S.T., 

and Chinchar, V.G., 2003, Diagnostic and molecular evaluation of three Iridovirus-associated salamander mortality 

events: Journal of Wildlife Diseases, v. 39, p. 556–566. 

Hoverman, J.T., Gray, M.J., Miller, D.L., and Haislip, N.A., 

2012, Widespread occurrence of ranavirus in pond-breeding 

amphibian populations: EcoHealth, v. 9, no. 1, p. 36–48, 

http://dx.doi.org/ 10.1007/s10393-011-0731-9.

Hyatt, A.D., Gould, A.R., Zupanovic, Z., Cunningham, A.A., 

Hengstberger, S., Whittington, R.J., Kattenbelt, J., and 

Coupar, B.E.H., 2000, Comparative studies of piscine and 

amphibian iridoviruses: Archives of Virology, v. 145, p. 

301–331. 

Miller, D. L., Gray, M. J., and Storfer, A., 2011, Ecopathology 

of ranaviruses infecting amphibians: Viruses, v. 3, no. 11, p. 

2351–2373, accessed August 30, 2012, at http://fwf.ag.utk.

edu/mgray/Publications/Milleretal2011.pdf.
Appendix 1 33

Appendix 1
34 U.S. Geological Survey, National Wildlife Health Center, 2011 Annual Report of Selected Wildlife Diseases

Table A1. Total number of bird disease cases by bird group and proportion of bird disease cases by bird group.

Total number of disease cases

Group Common 

name

Disease

Botulism 

C

Botulism 

E

Avian 

cholera

West 

Nile 

virus

SalmonellosisVirulent 

Newcastle

Lead 

poisoning

Trematode

 infection Total

Anseriformes Ducks, geese 

and swans

267 50 151 50 44 45 92 95 794

CharadriiformesShorebirds, 

gulls and 

terns 

155 47 33 45 54 29 29 29 421

Ciconiiformes Wading birds 

herons and 

egrets

33 9 9 7 15 7 7 7 94

ColumbiformesDoves and 

pigeons 

1 1 1 1 2 1 1 1 9

Falconiformes Falcons, 

hawks and 

eagles

15 14 18 33 20 14 35 14 163

Galliformes Upland 

gamebirds

1 1 1 4 1 1 1 1 11

Gaviiformes Loons 3 12 3 3 3 3 9 3 39

Gruiformes Cranes, rails 

and coots

11 3 13 2 2 2 2 9 44

Passeriformes Songbirds 17 17 22 30 53 17 18 17 191

Pelecaniformes Pelicans and 

cormorants

53 15 7 15 15 7 4 4 120

PodicipediformesGrebes 23 14 10 7 7 6 6 6 79

Strigiformes Owls 3 3 4 9 3 3 3 3 31
Appendix 1 35

Table A1. Total number of bird disease cases by bird group and proportion of bird disease cases by bird group.—Continued

Proportion of disease cases

Disease

Group Common 

name Botulism 

C

Botulism 

E

Avian 

cholera

West 

Nile 

virus

SalmonellosisVirulent 

Newcastle

Lead 

poisoning

Trematode 

 infection

Anseriformes Ducks, geese 

and swans

33.63 6.30 19.02 6.30 5.54 5.67 11.59 11.96

CharadriiformesShorebirds, 

gulls and 

terns

36.82 11.16 7.84 10.69 12.83 6.89 6.89 6.89

Ciconiiformes Wading birds 35.11 9.57 9.57 7.45 15.96 7.45 7.45 7.45

ColumbiformesDoves and 

pigeons

11.11 11.11 11.11 11.11 22.22 11.11 11.11 11.11

Falconiformes Falcons, hawks 

and eagles

9.20 8.59 11.04 20.25 12.27 8.59 21.47 8.59

Galliformes Upland 

gamebirds

9.09 9.09 9.09 36.36 9.09 9.09 9.09 9.09

Gaviiformes Loons 7.69 30.77 7.69 7.69 7.69 7.69 23.08 7.69

Gruiformes Cranes, rails 

and coots

25.00 6.82 29.55 4.55 4.55 4.55 4.55 20.45

Passeriformes Songbirds 8.90 8.90 11.52 15.71 27.75 8.90 9.42 8.90

Pelecaniformes Pelicans and 

cormorants

44.17 12.50 5.83 12.50 12.50 5.83 3.33 3.33

PodicipediformesGrebes 29.11 17.72 12.66 8.86 8.86 7.59 7.59 7.59

Strigiformes Owls 9.68 9.68 12.90 29.03 9.68 9.68 9.68 9.68
36 U.S. Geological Survey, National Wildlife Health Center, 2011 Report of Selected Wildlife Diseases Table A2. Disease cases, by State, from 2001 to 2005, and from 2006 to 2011. State Botulism 

C

Botulism 

E

Avian

Cholera Salmonellosis

West 

Nile

Virus

Trematode Lead Poisoning Virulent Newcastle Ranavirus Chytridio- mycosis Total disease events Amphibian disease events Avian disease events

Cases from 2001 to 2005

Alabama 0 0 0 0 0 0 0 0 0 0 0 0 0

Alaska 0 0 0 3 0 0 6 0 2 0 11 2 9

Arizona 0 0 0 1 0 0 13 0 1 1 16 2 14

Arkansas 0 0 0 0 0 0 12 0 0 0 12 0 12

California 23 0 24 3 2 0 5 0 4 4 65 8 57

Colorado 1 0 0 0 0 0 8 0 0 0 9 0 9

Connecticut 0 0 0 0 0 0 0 0 0 0 0 0 0

District of 

Columbia 0 0 0 0 1 0 0 0 0 0 1 0 1

Delaware 0 0 0 0 0 0 0 0 0 0 0 0 0

Florida 1 0 1 2 2 0 8 0 1 0 15 1 14

Georgia 0 0 0 1 0 0 0 0 0 0 1 0 1

Hawaii 28 0 0 1 0 0 0 0 0 0 29 0 29

Idaho 2 0 1 0 0 0 11 0 0 0 14 0 14

Illinois 2 0 0 1 2 0 4 0 0 0 9 0 9

Indiana 1 0 0 0 0 0 0 0 0 0 1 0 1

Iowa 1 0 1 1 5 1 8 0 0 0 17 0 17

Kansas 1 0 0 1 1 0 1 0 0 0 4 0 4

Kentucky 1 0 0 0 2 0 1 0 1 0 5 1 4

Louisiana 0 0 1 1 0 0 1 0 0 0 3 0 3

Maine 0 1 0 0 0 0 5 0 3 1 10 4 6

Maryland 4 0 0 0 3 0 3 0 0 0 10 0 10

Massachusetts 2 0 0 2 0 0 0 0 0 0 4 0 4

Michigan 0 0 0 2 1 0 0 0 0 0 3 0 3

Minnesota 4 1 1 1 5 0 9 0 1 0 22 1 21

Mississippi 0 0 0 0 0 0 4 0 0 0 4 0 4

Missouri 0 0 4 1 2 0 9 0 0 0 16 0 16

Montana 6 0 0 0 3 0 4 0 1 2 16 3 13

Nebraska 1 0 5 1 1 0 11 0 0 0 19 0 19
Appendix 1 37

Table A2. Disease cases, by State, from 2001 to 2005, and from 2006 to 2011.—Continued

State Botulism C Botulism E Avian Cholera Salmonellosis West Nile Virus Trematode Lead Poisoning Virulent Newcastle Ranavirus Chytridio- mycosis Total disease events Amphibian disease events Avian disease events

Cases from 2001 to 2005

Nevada 2 0 0 1 1 0 0 1 0 2 7 2 5

New Hampshire 0 0 0 0 0 0 2 0 0 0 2 0 2New Jersey 0 0 0 0 0 0 1 0 0 0 1 0 1New Mexico 1 0 0 1 1 0 3 0 1 2 9 3 6New York 1 2 0 0 1 0 1 0 0 0 5 0 5North Carolina 0 0 0 0 0 0 1 0 1 0 2 1 1North Dakota 16 0 0 4 10 0 4 0 1 0 35 1 34

Ohio 2 0 0 0 4 0 1 0 0 0 7 0 7

Oklahoma 0 0 0 1 2 0 0 0 0 0 3 0 3

Oregon 2 0 0 0 0 0 3 0 1 2 8 3 5

Pennsylvania 1 2 0 0 0 0 2 0 0 0 5 0 5

Puerto Rico 1 0 0 0 0 0 0 0 0 0 1 0 1

Rhode Island 0 0 0 0 0 0 0 0 3 0 3 3 0

South Carolina 0 0 0 0 0 0 1 0 0 0 1 0 1

South Dakota 5 0 1 0 2 0 2 0 0 0 10 0 10

Tennessee 0 0 0 1 0 0 0 0 0 0 1 0 1

Texas 0 0 2 3 1 0 1 0 1 0 8 1 7

Utah 4 0 1 2 0 0 3 0 1 3 14 4 10Vermont 0 0 0 0 1 0 1 0 0 0 2 0 2Virginia 6 0 0 0 0 0 2 0 1 1 10 2 8Washington 1 0 0 1 0 0 2 0 0 0 4 0 4West Virginia 0 0 0 0 0 0 4 0 0 0 4 0 4Wisconsin 15 4 0 1 2 26 10 1 0 1 60 1 59Wyoming 0 0 0 1 0 0 3 0 3 0 7 3 4
38 U.S. Geological Survey, National Wildlife Health Center, 2011 Report of Selected Wildlife Diseases TTable A2. Disease cases, by State, from 2001 to 2005, and from 2006 to 2011.—Continued State Botulism 

C

Botulism 

E

Avian

Cholera Salmonellosis

West

Nile

Virus

Trematode Lead Poisoning Virulent Newcastle Ranavirus Chytridiomycosis Total disease events Amphibian disease events Avian disease events

Cases from 2006 to 2011

Alabama 0 0 0 1 0 0 0 0 0 0 1 0 1

Alaska 0 0 0 0 0 0 4 0 1 1 6 2 4

Arizona 3 0 0 0 1 0 5 0 0 1 10 1 9

Arkansas 0 0 3 1 0 0 3 0 0 1 8 1 7

California 19 0 27 1 3 0 2 2 2 3 59 5 54

Colorado 0 0 0 0 0 0 2 0 0 1 3 1 2

Connecticut 0 0 0 0 0 0 1 0 0 0 1 0 1

Delaware 1 0 0 0 0 0 1 0 0 0 2 0 2

District of 

Columbia

2 0 0 0 0 0 0 0 0 0 2 0 2

Florida 12 0 0 1 0 0 0 0 1 0 14 1 13

Hawaii 33 0 0 0 0 0 0 0 0 0 33 0 33

Idaho 3 1 1 0 1 0 1 0 3 0 10 3 7

Illinois 2 0 1 0 0 0 3 0 0 1 7 1 6

Indiana 1 0 0 1 0 0 1 0 0 0 3 0 3

Iowa 0 0 2 0 0 0 2 0 0 0 4 0 4

Kansas 1 0 3 1 0 0 1 0 0 0 6 0 6

Kentucky 1 0 0 0 0 0 0 0 0 0 1 0 1

Louisianna 0 0 1 0 0 0 4 0 0 0 5 0 5

Maine 0 0 0 2 0 0 0 0 1 0 3 1 2

Maryland 5 0 0 3 1 0 0 1 6 0 16 6 10

Massachusettes 1 0 0 5 0 0 0 0 1 0 7 1 6

Michigan 0 13 0 0 0 0 1 0 0 0 14 0 14

Minnesota 7 0 2 7 6 19 8 10 0 1 60 1 59

Mississippi 0 0 1 0 0 0 0 0 0 0 1 0 1

Missouri 0 0 7 1 1 0 1 0 0 0 10 0 10

Montana 10 1 0 3 7 8 1 0 0 0 30 0 30

Nebraska 0 0 5 0 2 0 4 0 1 0 12 1 11
Appendix 1 39

Table A2. Disease cases, by State, from 2001 to 2005, and from 2006 to 2011. —Continued

State Botulism C Botulism E Avian Cholera Salmonellosis West Nile Virus Trematode Lead Poisoning Virulent Newcastle Ranavirus Chytridiomycosis Total disease events Amphibian disease events Avian disease events

Cases from 2006 to 2011

Nevada 5 0 0 0 1 0 0 0 0 0 6 0 6

New Hampshire 0 0 0 0 0 0 2 0 0 0 2 0 2

New Jersey 0 0 0 2 0 0 0 0 0 0 2 0 2

New Mexcio 3 0 0 0 0 0 0 0 0 0 3 0 3

New York 0 4 0 0 0 2 0 0 1 4 11 5 6

North Carolina 0 0 0 1 0 0 0 0 0 1 2 1 1

North Dakota 20 2 7 16 8 0 0 2 0 0 55 0 55

Ohio 1 0 0 0 0 0 3 0 0 0 4 0 4

Oklahoma 0 0 1 0 0 0 1 0 0 0 2 0 2

Oregon 1 0 5 1 3 0 11 0 1 2 24 3 21

Pennsylvania 3 2 0 0 0 0 2 0 1 0 8 1 7

Puerto Rico 1 0 0 0 0 0 0 0 0 0 1 0 1

Rhode Island 0 0 0 0 0 0 0 0 3 0 3 3 0

South Dakota 10 0 0 0 5 0 2 1 0 0 18 0 18

Tennessee 0 0 0 0 0 0 1 0 1 0 2 1 1

Texas 2 0 3 1 0 0 1 0 0 0 7 0 7

Utah 10 0 5 1 0 0 0 1 0 0 17 0 17

Vermont 0 0 0 1 0 0 0 0 0 0 1 0 1

Virginia 1 0 0 0 0 0 2 0 0 1 4 1 3

Washington 2 0 0 1 0 0 7 0 0 0 10 0 10

West Virginia 0 0 0 2 0 0 2 0 0 0 4 0 4

Wisconsin 7 7 0 2 3 25 5 5 2 0 56 2 54

Wyoming 0 0 1 1 0 0 1 0 4 0 7 4 3

Green and others—U.S. Geological Survey, National Wildlife Health Center, 2011 Report of Selected Wildlife Diseases—Scientific Investigations Report 2012–5271