| 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.
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 * www.cdc.gov/eid * 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
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 * www.cdc.gov/eid * 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 firstname.lastname@example.org
Corvidae Feather Pulp and West Nile Virus Detection
Emerging Infectious Diseases ** Vol. 10, No. 5, May 2004 909