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West Nile Virus Feather Pulp Detection
 Corvidae Feather Pulp and West Nile Virus Detection

Douglas E. Docherty,* Renee Romaine Long,* Kathryn M. Griffin,* and Emi K. Saito*

We evaluated cloacal swab, vascular pulp of flight

feather, and kidney and spleen pool samples from carcasses

of members of the family Corvidae as sources of West

Nile virus (WNV). The cloacal swab, kidney and spleen

pool, and feather pulp were the source of WNV in 38%,

43%, and 77%, respectively, of the carcasses.

Samples from carcasses of the family Corvidae have

been used in the surveillance for West Nile virus

(WNV) since the virus was detected in the United States in

1999 (1). Various laboratories have used organs such as

brain, kidney, and spleen to isolate the virus in cell culture,

or to detect the virus by using a variety of techniques, or

both (2). WNV surveillance efforts (3,4) have reported

50%-70% of corvids as WNV positive. Postmortem oral

and cloacal swabs, along with the brain, are suitable for

detecting the virus in experimentally infected birds (5).

Previous studies have shown that WNV may be isolated

from feather pulp of experimentally infected crows

(National Wildlife Health Center, unpub. data). We

describe the results of WNV isolation attempts from cloacal

swabs, kidney and spleen pools, and the flight feathers

containing vascular pulp (6) of dead American Crows

(Corvus brachyrhynchos) and Blue Jays (Cyanocitta

cristata) that were found in the field and suspected of

being WNV infected.

The Study

Specimens were obtained at necropsy from a group of

28 American Crow and 56 Blue Jay carcasses. These birds

were submitted, from August through October, as field

cases in the course of the larger 2002 wildlife surveillance

effort for WNV at the U.S. Geological Survey (USGS),

National Wildlife Health Center, in Madison, Wisconsin.

Birds received for WNV surveillance were evaluated for

WNV but not for cause of death. Kidney and spleen pools,

cloacal swab, and feather pulp were collected from each of

these 84 birds received from the following nine states:

Alabama, Illinois, Kansas, Maryland, Missouri, North

Dakota, Pennsylvania, Texas, and Virginia. The birds in

this study were obtained after WNV was initially detected

in birds from each state; only those carcasses judged to be

fresh were sampled.

At necropsy each bird was examined for wing flight

feathers (remiges) and tail flight feathers (retrices) that

contained vascular pulp. These feathers were pulled from

the feather follicle and aseptically cut at the distal end of

the umbilicus (6). The umbilicus, containing vascular pulp,

was placed in viral transport media (7). At least one and up

to three feathers were collected from American Crows and

at least four and up to six were collected from Blue Jays.

Also at necropsy, cloacal samples were taken with Dacron

swabs that were swirled in viral transport media, squeezed

out against the side of the collection tube, and then discarded.

Kidney and spleen samples were aseptically collected

at necropsy. Samples were kept chilled at 4°C, and

any not processed within 24 hours after they were obtained

were stored at -80°C.

In the laboratory, a 10% (wt/volume) kidney and spleen

pool suspension was prepared in viral transport media. The

kidney and spleen pool suspension was blended in a

Stomacher 400 Circulator (Seward, Norfolk, UK) until it

appeared homogeneous. The vascular pulp was aseptically

removed from the umbilicus with forceps, and viral transport

media was added to the available feather pulp mass to

produce a 10% (wt/volume) suspension. The cloacal swab

and feather pulp suspensions were vortexed until they

appeared homogeneous. The kidney and spleen pool, cloacal

swab, and feather pulp suspensions were centrifuged at

800 x g for 30 min at 4°C, and 1 mL of the supernatant was

injected onto an established Vero (ATCC CRL-1587) cell

monolayer in 12-cm2 (culture surface area) bottles. These

bottles were incubated at 37°C and 2% CO2 and read periodically

over 7 days for viral cytopathic effect (CPE). To

screen for WNV, cell culture bottles showing viral CPE

involving at least 75% of the Vero monolayer were harvested

after one freeze-and-thaw cycle and tested for

WNV by reverse transcriptase-polymerase chain reaction

(RT-PCR) (8). The RT-PCR test was also used to determine

whether feather pulp or cloacal swab suspensions, negative

for virus isolation, contained quantities of WNV below the

detection level of cell culture. The number of WNV plaque

forming units (PFU) was determined (9) to evaluate the

quantity of virus in various kidney and spleen pool, cloacal

swab, and feather pulp samples.

With the screening method described here, WNV was

isolated from 65 (77%) of 84 corvids. Of the 65 WNV-positive

birds, WNV was isolated from 100% (65/65) of the

feather pulp samples, 55% (36/65) of the kidney and

spleen pool samples, and 49% (32/65) of the cloacal

swabs. WNV was isolated from all three samples for 25%

(21/84) of all birds tested. Attempts at virus isolation were

significantly (p < 0.001) more successful from feather pulp

than from either kidney and spleen pool or cloacal swab.

Emerging Infectious Diseases * * Vol. 10, No. 5, May 2004 907

*National Wildlife Health Center, Madison, Wisconsin, USA

The ability to successfully isolate WNV from either the

kidney and spleen pool or the cloacal swab was essentially

the same (p > 0.5).

The feather pulp or cloacal swab samples from birds

from which WNV was not isolated were also negative for

WNV by using RT-PCR. Comparisons of the quantity of

WNV in 0.1 mL of sample suspension indicated that more

PFU of WNV were in the feather pulp than cloacal swab or

kidney and spleen pool suspensions (Table). A comparison

of the number of PFU in the feather pulp samples and cloacal

swabs from the same 12 birds showed that the feather

pulp had significantly (p < 0.0005) more.


WNV isolation from feather pulp is a relatively sensitive

assay for surveillance of corvid carcasses. Of the 84

tested, 77% (65/84) of the birds were WNV positive by

feather pulp alone, 43% (36/84) were positive by kidney

and spleen pool alone, and 38% (32/84) were positive by

cloacal swab alone. On the basis of our determination of

the WNV titer in feather pulp, cloacal swab, and kidney

and spleen pool, these results could be explained by the

fact that much more virus was present in the feather pulp

suspension. The 23% (19/84) negative birds consisted of

16 Blue Jays and 3 American Crows, representing birds

that may have died of causes other than WNV infection.

Other causes may include other infectious agents, toxins,

or trauma not related to concurrent WNV infection.

Previous studies of Eastern equine encephalitis virus in

Ring-necked Pheasants (Phasianus colchicus) and avian

leucosis virus in domestic chickens (Gallus gallus) found

virus in feather pulp up to 7 days beyond detection in

blood (10-12). The virus titer in feather pulp was also

much greater than in blood or cloacal swab. Avian leukosis

virus could be detected in feather pulp even after antibody

was detected. In a recent publication (13), RT-PCR

was used to detect WNV in the "skin including feather

tips" of goslings experimentally infected with the virus.

The authors of that publication concluded that blood and

skin containing feather tips could, through cannibalism,

horizontally transmit sufficient virus to directly infect contact

control goslings.

The duration and timing of the molt is a limiting factor

in using the feather pulp sample. In much of the United

States, the American Crow molt will occur from July

through September, the Blue Jay from June through

October, and the Common Raven (Corvus corax) from

May through October (14). A feather pulp sample from

corvids in the United States will therefore be available during

the height of the WNV season. We recommend collecting

and testing the feather pulp, considering the apparent

high rate of success in detecting WNV.

The feather pulp sample is nonlethal and could be taken

from birds trapped live, sampled, and released. Sufficient

WNV appears to be available in samples obtained from

corvid carcasses suspected to be WNV infected to infect

cell culture. since none of the negative cloacal swab or

feather pulp samples were positive by RT-PCR. However,

for subclinical infections or from other species of birds,

additional testing may be necessary to determine whether

the amount of virus available in feather pulp will be sufficient

for detection by virus isolation or RT-PCR.


The authors thank Nathan Ramsay, Dottie Johnson, Heather

Gutzman, and Michelle Oates for obtaining the postmortem specimens

used in this study and Scott Wright, Carol Meteyer, Robert

McLean, and Tracy McNamara for various contributions.

Funding was provided by the U.S. Geologic Survey and the

Centers for Disease Control and Prevention.

Mr. Docherty is diagnostic virologist for the Department of

the Interior, U.S. Geological Survey, National Wildlife Health

Center. His primary responsibilities are wildlife virology diagnostics

and wildlife disease research projects.


1. Eidson M, Komar N, Sorhage F, Nelson R, Talbot T, Mostashari F, et

al. Crow deaths as a sentinel surveillance system for West Nile virus

in the Northeastern United States, 1999. Emerg Infect Dis


2. Marfin A, Petersen L, Eidson M, Miller J, Hadler J, Farello C, et al.

Widespread West Nile virus activity, Eastern United States, 2000.

Emerg Infect Dis 2001;7:730-5.

3. Hadler J, Nelson R, McCarthy T, Andreadis T, Lis M, French R, et al.

West Nile virus surveillance in Connecticut in 2000: an intense epizootic

without high risk for severe human disease. Emerg Infect Dis


4. Pannela N, Kerst A, Lanciotti R, Brant P, Wolf B, Komar N.

Comparative West Nile virus detection in organs of naturally infected

American crows. Emerg Infect Dis 2001;7:754-5.

5. Komar N, Lanciotti R, Bowen R, Langevin S, Bunning M. Detection

of West Nile virus in oral and cloacal swabs collected from bird carcasses.

Emerg Infect Dis 2002;8:741-2.

6. McKibben JS, Harrison GJ. Clinical anatomy. In: Harrison GJ,

Harrison LR, editors. Clinical avian medicine and surgery.

Philadelphia: WB Saunders Co.; 1986. p. 32-7.

7. Docherty D, Slota P. Use of Muscovy duck embryo fibroblasts for the

isolation of viruses from wild birds. J Tissue Cult Methods



908 Emerging Infectious Diseases * * Vol. 10, No. 5, May 2004

Table. Logarithmic titers of West Nile virus infectious particles

(per 0.1 mL of 10% tissue suspension) present in each type of

sample, as detected by plaque assay in Vero cells

Type of sample No. tested Median (range)

Kidney/spleen pool 7 1.0 (<1.0 to 3.3)

Cloacal swab 12 1.9 (<1.0 to 4.0)

Vascular pulp of flight feather 12 4.9 (3.5 to >7.4)

8. Lanciotti R, Kerst A, Nasci R, Godsey M, Mitchell C, Savage H, et

al. Rapid detection of West Nile virus from human clinical specimens,

field-collected mosquitoes, and avian samples by a TaqMan

reverse transcriptase-PCR assay. J Clin Microbiol 2000;38:4066-71.

9. Beaty B, Calisher C, Shope R. Arboviruses. In: Lennette EH,

Lennette DA, Lennette ET, editors. Diagnostic procedures for viral,

rickettsial, and chlamydial infections; 7th ed. Washington: American

Public Health Association; 1995. p. 204-5.

10. Satriano S, Luginbuhl R, Wallis R, Jungherr E, Williamson L.

Investigation of Eastern equine encephalomyelitis IV: susceptibility

and transmission studies with virus of pheasant origin. Am J Hygiene


11. Zavala G, Jackwood M, Hilt D. Polymerase chain reaction for detection

of avian leucosis virus subgroup J in feather pulp. Avian Dis


12. Sung H, Reddy S, Fadly A. High virus titer in feather pulp of chickens

infected with subgroup J avian leucosis virus. Avian Dis


13. Banet-Noach C, Simanov L, Malkinson M. Direct (non-vector) transmission

of West Nile virus in geese. Avian Path 2003;32:489-94.

14. Pyle P. Identification guide to North American birds. Bolinas (CA):

Slate Creek Press; 1997. p. 297-314.

Address for correspondence: Douglas Docherty, National Wildlife Health

Center, 6006 Schroeder Rd., Madison, WI 53711, USA; fax: 608-270-

2415; email

Corvidae Feather Pulp and West Nile Virus Detection

Emerging Infectious Diseases ** Vol. 10, No. 5, May 2004 909


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