|UFDC Home||myUFDC Home | Help|
This item has the following downloads:
EPIZOOTIOLOGY OF FELINE LEUKEMIA VIRUS IN THE FLORIDA PANTHER
MARK WILLIAM CUNNINGHAM
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
Mark William Cunningham
"The role of disease in wildlife conservation has probably been radically underestimated"
(Aldo Leopold, 1933).
Most of the information presented in this thesis would not be possible without the
expertise of houndsman Roy McBride who began capturing panthers in the 1970s. Also
very much appreciated are the efforts of the biologists on the Florida Fish and Wildlife
Conservation Commission (FWC) panther capture team including Darrell Land, David
Shindle, and Mark Lotz. Veterinarians with FWC who collected samples or data used in
this study include Drs. Melody Roelke, Mike Dunbar, Sharon Taylor, Dave Rotstein, and
Kristin Mansfield. Researchers with the National Park Service also collected samples and
include Deborah Jansen, Steve Schultz, and Dr. Emmett Blankenship.
Veterinary pathologists have been especially helpful with this study and include
Drs. Scott Terrell, Claus Buergelt, and Bruce Homer.
I would especially like to thank my advisor Dr. Donald Forrester for his patience
and guidance. I also appreciate the guidance of other committee members including Drs.
Julie Levy, Mel Sunquist, and Rick Alleman.
I would also like to thank collaborators on this project including Drs. Meredith
Brown, Stephen J. O'Brien, and Warren Johnson at the National Cancer Institute; and
Drs. Kathleen Hayes and Lawrence Mathes at the Ohio State University. I greatly
appreciate the advice and support provided by Drs. Scott Citino, Cynda Crawford, and
William Hardy, Jr. Finally, I am indebted to Richard Kiltie for assistance with statistical
This project was fully funded by the Florida Fish and Wildlife Conservation
Commission through the Federal Endangered Species Project E-1 and the Florida Panther
Restoration and Management Trust Fund.
TABLE OF CONTENTS
A C K N O W L E D G M E N T S ................................................................................................. iv
LIST OF TABLES .................................................... ............ ............. .. viii
LIST OF FIGURES ......... ......................... ...... ........ ............ ix
1 IN TR OD U CTION ............................................... .. ......................... ..
B background ............................................................... .. ........ ...............
The Florida Panther ................................................. .... ...... .............. ..
Feline Leukemia Virus ................................... ...... ...........4....4
O b j e c tiv e s ........................................................................................................1 1
2 M ATERIALS AND M ETHOD S ........................................ ......................... 13
Study Area and Period .............. .. .......... ...................... ................. 13
Florida Panther Capture and Immobilization .................................. ............... 13
Physical Exam nation ...... ............................... ......... .... .. ................14
Live-capture Sample Collection ...... ..................... .................14
V accination and Treatm ent........................................... ........................... 15
R adio-instrum entation .............. ..................... ................... ............... 16
N e o n ata l K itten s ................................................................................................... 16
N e c ro p sy .......................................................................................................1 6
Specim en Storage ................... .................................................. ...... . ......... 17
Age D term nation and Genetic Status.................................. ....................... 17
D diagnostics ................... .................................................... . ........... 17
Enzyme-linked Immunosorbent Assay Antibody ............................................17
Enzyme-linked Immunosorbent Assay Antigen...............................................17
Immunofluorescent Assay and Immunohistochemistry ....................................18
Polymerase Chain Reaction, Genetic Sequencing, and Viral Culture.................18
Complete Blood Count and Serum Chemistry ..................................................19
O their D iagnostic Testing.......................................................... ............... 19
S ta tistic s .............................................. .. .................... ................ 1 9
3 R E SU L T S ....................................................... 22
D diagnostic T ests...................... .... .................................... ............... ... ............ 22
Enzyme-linked Immunosorbent Assay Antibody ............................................22
Enzyme-linked Immunosorbent Assay Antigen ..............................................23
Immunofluorescent Assay and Immunohistochemistry ....................................24
O th er sero lo g y ................................................... ................ 2 4
C lin ical F in din g s................................................. ................ 2 5
C clinical P ath ology .............................. ......................... ... ...... .. .... ............2 5
P ath o lo g y ...................................................................................................2 5
G r o s s ....................................................................................................... 2 5
M ic ro sc o p ic ............................................................................. 2 6
O opportunistic infections ........................................ .......................... 26
M o reality .................................................................................. 2 6
4 DISCUSSION ....................................................... ........... .... .......... 28
D ia g n o stic s ........................................................................................................... 2 8
E p iz o o tio lo g y ............. ......... .. .... ........... .................................................. 3 0
H history of E xposure.......... ..... .................................................... .. .......... .... 30
Prevalence and D distribution ........................................ .......................... 31
Outcome Following Exposure.................................... ..................... 31
Self-lim iting infections...................... .. .. .......... .. ................. ............... 32
P ersistent infection s.......... ............................................ ........ .... ......... 33
Epizootiology ................................... .......... ......................... 38
C on clu sion ...................... ................ ...................................................... 4 0
Further Research ............... .............. ............... ........ ...... ..............41
A FLORIDA PANTHER/TEXAS PUMAS SAMPLED DURING THE STUDY
P E R IO D ...................................... ......................................................42
B CASE REPORTS: ANTIGENEMIC FLORIDA PANTHERS..............................49
F P 1 1 5 ....................................................................... 4 9
F P 1 0 9 ....................................................................... 5 1
F P 1 2 2 ....................................................................... 5 2
F P 12 3 ...................................... ......... ..................... ................ 5 4
F P 1 3 2 ....................................................................... 5 5
L IST O F R E F E R E N C E S ......... .................................................................. ............... 60
B IO G R A PH IC A L SK E TCH ..................................................................... ..................70
LIST OF TABLES
A-i Florida panthers and Texas pumas tested for feline leukemia virus (FeLV)
antigen by ELISA 1 July 2002, to 5 June 2005..................... .............. ............... 43
B-1 Selected hematological and serum biochemical values for Florida panthers
testing positive for feline leukemia virus (FeLV) antigen by ELISA 1 July 2002
to 5 June 2005. ..................................................... ................. 59
LIST OF FIGURES
1-1 Outcome following exposure to feline leukemia virus in domestic cats..................12
2-1 Study area in south Florida, USA ... ..................... .... ..... ................ 21
3-1 Distribution of positive feline leukemia virus positive ELISA antibody optical
densities in Florida panthers/Texas pumas by region and year 1990-2005 ...........23
3-2 Feline leukemia virus (FeLV) ELISA antigen results for panthers >1 yr, not
previously FeLV vaccinated, and sampled in South Forida between 1 July 2002
an d 5 Ju n e 2 0 0 5 .................................................................... 2 7
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science
EPIZOOTIOLOGY OF FELINE LEUKEMIA VIRUS IN THE FLORIDA PANTHER
Mark William Cunningham
Chair: Donald J. Forrester
Major Department: Wildlife Ecology and Conservation
Feline leukemia virus (FeLV) has been reported only rarely in non-domestic felids
and was not detected in Florida panthers (Puma concolor coryi) during almost 20 yr of
routine surveillance. The finding of two FeLV antigen-positive panthers during the 2002-
2003 capture season led to a prospective and retrospective investigation of the
epizootiology of this disease in the population. Archived serum was tested for FeLV
antibodies to assess history of exposure. To determine prevalence and distribution,
panthers were captured throughout their range and tested for FeLV antigen by ELISA.
Positive tests were confirmed by immunofluorescent antibody (IFA) test and viral
culture. The outcome following exposure in panthers was inferred from ELISA antigen
and antibody, IFA, and PCR results. All infected panthers were monitored by radio-
telemetry and necropsied following detection of a mortality signal. Between 1990 and
2005, the prevalence of positive antibody tests increased significantly and were
concentrated in the northern portion of panther range. The prevalence of antigenemia
(positive ELISA antigen) among panthers and Texas pumas >1 yr of age, not previously
vaccinated for FeLV, and sampled between July 2002 and June 2005, was 7% (5 of 71).
Antigenemic panthers were captured or recovered in the Okaloacoochee Slough State
Forest (OKS) in the northern portion of panther range. All antigenemic panthers were
positive by viral culture and three were IFA positive at capture. Clinical signs and clinical
pathology at capture (n = 4) included lymphadenopathy, moderate to severe anemia,
lymphopenia, and acute lymphoblastic leukemia. All infected panthers died during the
study period; causes of deaths were septicemia (n = 2), intraspecific aggression (n = 2),
and unknown (n = 1). Average time from diagnosis to death was 9.25 (SD +10.3) wk in
antigenemic panthers. Following exposure, panthers developed transient, latent, or
persistent infections. The high localized prevalence of antigenemic panthers in OKS
(45.5%) demonstrates the potential impact of this disease on the population. Management
to control the epizootic currently includes vaccination and test-removal. No new cases
have been diagnosed since July 2004.
The Florida panther (Puma concolor coryi) is one of the most endangered mammals
in North America, at one time numbering as few as 30 individuals. With protection and
management the population has rebounded to almost 100; however, the panther is now
threatened by feline leukemia virus (FeLV). Feline leukemia virus infection is a fatal
infectious disease, common to domestic cats (Felis catus), that is rare in non-domestic
felids. Routine FeLV antigen testing in panthers was negative for almost 20 yr until two
positive panthers were detected during the 2002-2003 capture season. These findings
resulted in a prospective and retrospective investigation into the epizootiology of this
disease in the panther population. Information gained from this research will not only be
used to help manage the epizootic in this critically endangered population but may also
benefit managers of other non-domestic felid populations.
The Florida Panther
The Florida panther is an endangered subspecies of puma whose range was once
contiguous with other puma subspecies including the Texas puma (P. concolor
stanlyana). By the early part of the 20th Century; however, habitat destruction,
exploitation, and human population growth had reduced the panther to an isolated
remnant population. The panther was eliminated eventually from all previous range with
the exception of the relatively inaccessible and, historically, undesirable Big Cypress and
Everglades ecosystems of south Florida. Protection of the panther began with state
classification as a game animal in 1950 followed by complete state protection in 1958.
The panther was listed federally as an endangered species in 1967. Nevertheless, the
population dwindled to an estimated 20 to 30 individuals by the early 1970's (Nowak and
Researchers noticed morphologic differences among panthers from different areas
of south Florida. Subsequent genetic analyses revealed two genotypes: 1) original or
canonical Florida panthers, concentrated in the Big Cypress ecosystem, and 2) Florida
panther/South American puma intercrosses which primarily occupied the Everglades
ecosystem (O'Brien et al., 1990). The canonical genotype traced its lineage from the
original remnant population while the South American puma intercrosses likely resulted
from the release of Florida panther/captive puma hybrids into the free-ranging panther
population between 1957 and 1967 (Vanas, 1976). Panthers with genetic evidence of
South American puma ancestry, although representing a minority, had a greater genetic
diversity and fewer congenital anomalies than panthers retaining the canonical genotype
(Roelke et al., 1993a). Among canonical panthers, the level of mitochondrial DNA
variation, frequency of polymorphic allozyme loci, and average heterozygocity of
allozyme loci was lower than any other similarly studied feline except the cheetah
(Acinonyx ubatus) (O'Brien et al., 1990; Newman et al., 1985; Roelke et al., 1993a).
The consequences of inbreeding in panthers were believed to have included
cryptorchidism (Roelke et al., 1993a; Mansfield and Land, 2002), atrial septal defects
(Cunningham et al., 1999), poor seminal traits (Barone et al., 1994), and poor fecundity
(Roelke et al., 1993a). Putative impaired immunocompetence was suspected to increase
susceptibility to parasites and infectious diseases including dermatophytosis (Rotstein et
al., 1999). Many of these traits are still seen in canonical Florida panthers today.
Without intervention the Florida panther was predicted to become extinct within 25
to 40 yr (Seal and Lacy, 1989). However, in 1995 eight female Texas pumas were
released into south Florida as part of a genetic restoration program (Seal, 1994). The
resultant introgression was designed to restore the genetic diversity to levels comparable
to other puma subspecies and to lower the incidence of congenital anomalies in the
As of September 2004, over half of the population had Texas puma genes (D. Land,
pers. commun.). The distribution of genotypes was not uniform however, with more
canonical panthers present in the northern portion of panther range. Recent microsatellite
DNA analyses also provided evidence for a third and more recent introgression. Several
captive pumas of unknown western ancestry escaped from the Seminole Indian
Reservation (SIR) north of Big Cypress National Preserve (BCNP) between 1996 and
1999. Although most were eventually recaptured, successful breeding with free-ranging
panthers apparently occurred, and evidence of this genotype was present in 6-10% of
panthers sampled between 2000 and 2004 (D. Land, pers. commun.). This genotype was
concentrated also in the northern portion of panther range.
The prevalence of congenital anomalies among intergrades was reduced greatly and
was limited to the occasional kinked tail or cowlick. As a result of the genetic
introgressions, both deliberate and unintentional, and other management measures, the
panther population had rebounded to a minimum of 87 by 2003 (McBride, 2003).
However, this increase in density may have resulted in an increased risk of infectious
disease transmission and expansion of the wildland-urban interface. These factors may
have set the stage for the current FeLV epizootic.
Feline Leukemia Virus
Feline leukemia virus is a Gammaretrovirus in the family Retroviridae. Following
penetration of the host cell by the viral RNA, reverse transcriptase transcribes viral RNA
into double-stranded DNA which is then incorporated into the host genome. Incorporated
viral DNA, known as provirus, codes for viral proteins and serves as a template for the
production of viral RNA. There are numerous strains of FeLV and few isolates in nature
are identical (Hoover and Mullins, 1991). Feline leukemia virus is classified into
subgroups A, B, and C based on envelope antigens (Jarrett et al., 1973; Sarma and Log;
1973). All FeLV-infected cats carry subgroup A (Jarrett et al., 1978), which is the least
pathogenic and only transmissible form. Subgroup C results from mutation of subgroup
A while subgroup B arises from recombination between subgroup A and endogenous
retroviral DNA (enFeLV) (reviewed by Miyazawa, 2002). EnFeLV are non-coding, non-
immunogenic sequences (Mandel et al., 1979; Rigby et al., 1992) that became
incorporated in the domestic cat genome early in their phylogenetic history. Most non-
domestic felids, including Florida panthers, do not have enFeLV.
The domestic cat is the definitive host for FeLV and the virus has a worldwide
distribution. Although several non-felid cell lines have shown in vitro susceptibility
(Nakata et al., 2003) infection has not been described in non-felid species. The worldwide
prevalence of FeLV in healthy domestic cat populations ranges from 1-8% (Levy, 1999)
with prevalences over 30% in some closed populations (Grant et al., 1980; Gertsmann,
1985). There is evidence that the overall prevalence of FeLV in domestic cats is
decreasing, possibly due to vaccination and other control measures (Levy and Crawford,
2005). In Florida, the prevalence among feral cats is less than 4% (Lee et al., 2002).
Infection is more prevalent among male cats, mixed breed cats, and cats between 1 and 7
yr of age (Levy, 2005). The highest infection rate occurred in cats less than 2 yr of age
(Levy, 1999). In contrast, the prevalence of FeLV antibodies, indicating exposure,
continues to increase with age (Rogerson et al., 1975).
Feline leukemia virus is an enveloped virus and is therefore quite fragile. The virus
immediately begins losing viability outside of the host and, on dry surfaces, is completely
inactivated between two and three hr (Francis et al., 1979). Therefore transmission is
primarily by direct contact. The virus is shed in highest concentrations in the saliva
(Francis et al., 1977), and horizontal transmission occurs primarily via the oronasal route
and by bite wounds. Prolonged contact is generally necessary for effective transmission
(Hardy et al., 1973). Transplacental and transmammary transmission of the virus are also
important (Hardy et al., 1976).
Following exposure most domestic cats will eventually clear the virus while
approximately one-third will become persistently infected and eventually succumb to
FeLV related diseases. However, there is a dynamic relationship between the host and
virus, and progression of disease depends on a number of factors. Outcome following
exposure depends on host age (Hoover et al., 1976), genetics (Hoover and Mullins,
1991), and immunocompetence (Hoover et al., 1980), as well as route of exposure, virus
burden, and strain of virus (Rojko and Kociba, 1991; Hoover and Mullins, 1991). The
progression of infection can be predicted by provirus burden using quantitative
polymerase chain reaction (PCR) (Hofmann-Lehmann et al., 2001). Cats clearing
infection early have no or low provirus burdens, those latently infected retain moderate
levels of provirus, while those becoming persistently infected have high provirus burdens
that peak at about 4 wk post-exposure (Hofmann-Lehmann et al., 2001). These
researchers also demonstrated an inverse correlation between ELISA antibodies and
provirus load beginning approximately 3 wk post-exposure. Cats that resisted persistent
infection had a more pronounced humoral response and lower provirus burdens than cats
that progressed to persistent infections. Cell-mediated immunity is important also in the
early immune response to FeLV infection (Flynn et al., 2002). Regardless of the
outcome, the course of infection is established usually by 8 wk post-exposure (Torres et
al., 2005). Outcome following exposure in domestic cats is summarized in Fig. 1-1.
Following exposure the virus replicates in local lymphoid tissues. Approximately
40% of cats mount an effective immune response and clear the virus before further
progression (Hoover and Mullins, 1991). These cats remain antigen and provirus negative
throughout their lives (Torres et al., 2005). If the infection progresses, however, viral
replication within a small number of circulating leukocytes will lead to infection of
lymphoid organs including the thymus, spleen, and lymph nodes (Rojko et al., 1979).
Cats at this stage may be transiently antigenemic and may even be briefly infectious.
Clinical signs during this primary viremia may include fever, lethargy, leukopenia,
anemia, and lymphadenopathy (Pedersen et al., 1990; Levy, 1999). However,
approximately 50% of cats reaching this stage are still able to mount an effective immune
response and clear the infection (Hoover and Mullins, 1991). Failure of viral containment
will lead to infection of the bone marrow, salivary glands, and other tissues between 3
and 13 wk. Nevertheless, an adequate immune response early in this process may rescue
the cat from persistent infection. These cats will retain provirus in peripheral and marrow
leukocytes for variable periods and are considered latently infected. Latently infected cats
do not shed virus and are not infective to other cats. Reactivation of latent infections
following stress is possible but becomes less likely >1 yr post-infection (Pedersen et al.,
1984). Generally, cats recovering from transient or latent infections are immune to re-
Progression to persistent infection occurs in approximately 35% of exposed cats
and is characterized by infection of the bone marrow and the development of
cytosuppressive and cytoproliferative diseases. Severity and type of disease in
persistently infected cats depends on host age (Hoover et al., 1976), concurrent feline
immunodeficiency virus (FIV) infection, and virus subgroup and strain. Following
establishment of a persistent infection, a period of dormancy ensues lasting weeks to
years during which few if any clinical signs are apparent. Eventually, persistent infections
result in any of three clinical syndromes: immunosuppression, anemia, and/or neoplasia.
Immunosuppression is believed to result in opportunistic infections. Co-infections
were the most frequent finding in FeLV infected cats examined at North American
veterinary schools (Levy, 1999). Anemia, whether primary or secondary, is the next most
common clinical finding in FeLV infected cats. Anemias are most commonly non-
regenerative and include pure red cell aplasia, red blood cell macrocytosis, erythemic
myelosis, bone marrow infiltration, and anemia of chronic disease (Hardy, 1980a).
Finally, hematopoietic neoplasms may also result from FeLV infection. Lymphoma is the
most common FeLV-related neoplastic disease; leukemias, myeloproliferative diseases,
and fibrosarcomas are also common (Hardy, 1980a). Mortality among persistently
infected cats is approximately 5-fold that of uninfected cats and 83% die within 3.5 yr
(McClelland et al., 1980).
Co-infections of FIV and FeLV are believed to work synergistically to result in
more severe disease (Grindem et al., 1989; Pedersen et al., 1990; Hofmann-Lehmann et
al., 1997). Beebe et al. (1994) suggested that immunosuppression caused by pre-existing
FeLV infection affected disease development upon subsequent FIV infection. Feline
immunodeficiency virus infected cats experimentally infected with FeLV had more
severe disease with a more rapid onset than cats infected with either virus alone. Further,
CD4+ T-lymphocytes were much more depressed in co-infected cats than cats infected
with either virus alone (Hoffmann-Lehmann et al., 1995). It is unknown whether the
order of infection (FeLV or FIV first followed by the other) is important in the clinical
outcome (Hofmann-Lehmann et al., 1997). Finally, virus/virus interactions such as the
formation of FeLV/FIV pseudotypes does not appear to be a mechanism of disease
potentiation (Beebe et al., 1994).
The outcome following introduction of FeLV into naive domestic cat populations
depends on population size, density, dispersal patterns, and spatial and social structure
(Fromont et al., 1998a,b; Fromont et al., 2003). Based on computer models, FeLV
becomes established in large natural domestic cat populations at a prevalence of between
0.8% and 12.4% depending on the parameters used (Fromont et al., 1998a,b) and reduces
population size by 3% (Courchamp et al., 1997) to 7% (Fromont et al., 1997). Inclusion
of FIV in Courchamp's et al. (1997) models more than doubled the population impact of
FeLV. Fromont et al. (1998a) also predicted that FeLV fails to become established in
small isolated populations numbering <100 individuals although extinction of the virus
may take several years.
Feline leukemia virus can be diagnosed and staged using a variety of techniques.
The enzyme-linked immunosorbent assay (ELISA) antigen test is the most common
screening method. The ELISA detects soluble p27 antigen in blood (Lutz et al., 1980a)
usually within 3 wk post-infection (Hofmann-Lehmann et al., 2001). Positive test results
may indicate transient or persistent infection and are an indicator of viremia.
Confirmation of positives is accomplished by immunofluorescent assay (IFA), which
detects p27 antigen within neutrophils and platelets of blood smears (Hardy et al., 1973).
A positive IFA test indicates infection of the bone marrow and usually indicates
persistent infection. Viral culture is highly specific and may be used to detect transient,
latent, or persistent infections and to identify subgroup. Polymerase chain reaction is a
highly sensitive and specific technique that has been used to detect integrated provirus or
free FeLV in formalin-fixed tissues, fresh tissues, bone marrow, and blood. Most
transient and persistent infections are detectable by PCR 1 wk post-infection and all are
detectable by 2 wk (Hofmann-Lehmann et al., 2001). Detection of FeLV antibodies helps
stage the disease, especially identifying previous transient infections, but has little
importance in diagnosis. Feline leukemia virus ELISA antibodies are most frequently
found in those groups clearing the infection (Lutz et al., 1980b). Finally, sequencing of
virus is used to identify strain and subgroup. Expected test results during various stages
of FeLV infection are summarized in Fig. 1-1.
Infection of non-domestic felids by FIV, also a retrovirus, is relatively common and
usually does not result in clinical signs. Approximately 28% of Florida panthers carry the
puma lentivirus strain of FIV (Olmstead et al., 1992) and pathology has not been
observed. In contrast to FIV, FeLV infections in non-domestic felids are quite rare. Feline
leukemia virus infection has been documented in a handful of captive non-domestic felids
including a bobcat (Lynx rufus) (Sleeman et al., 2001), puma (Meric, 1984), clouded
leopard (Neofelis nebulosa) (Citino, 1986), and several cheetahs (A. jubatus) (Briggs and
Ott, 1986; Marker et al., 2003). Feline leukemia virus has also been isolated from a
leopard cat (F. bengalensis) cell line (Rasheed and Gardner, 1981). In all cases, the
source of infection was believed to be infected domestic cats.
Despite extensive testing for FeLV in free-ranging felid populations (Rasheed and
Gardner, 1981; Mochizuki et al., 1990; Roelke et al., 1993b; Paul-Murphy et al., 1994;
Hofmann-Lehmann et al., 1996; Miyazawa et al., 1997; Osofsky et al., 1996; Biek et al.,
2002; Munson et al., 2004; Riley et al., 2004; Ryser-Degioris et al., 2005) published
reports of FeLV infection have been limited to a puma (P. concolor) in California (Jessup
et al., 1993) and a sand cat (F. margarita) in Saudi Arabia (Ostrowski et al., 2003).
Approximately 10 to 24% of European wildcats (F. sylvestris sylvestris) were also FeLV
positive (Daniels et al., 1999; Fromont et al., 2000), although interbreeding with domestic
cats occurs frequently in this subspecies (Daniels et al., 1998).
There have been reports of positive FeLV test results in free-ranging non-domestic
felids that were not confirmed with additional tests. Rickard and Foreyt (1992) detected
FeLV antigen in 2 of 2 free-ranging pumas found dead in Washington but virus isolation
was not attempted. Schmitt et al. (2003) diagnosed FeLV infection by IFA in 11 of 16
(69%) captive and free-ranging felids from Brazil, a biologically inconsistent percentage,
but did not confirm the results by ELISA antigen or viral culture.
Testing for FeLV antibodies has been performed only rarely on samples collected
from non-domestic felids. Feline leukemia virus antibodies were found in a transiently
infected captive clouded leopard (Citino, 1986) and two captive Siberian tigers (Panthera
tigris altaica) (Meric, 1984).
Most infections in non-domestic felids were self-limiting. In a survey of North
American zoos, 7 of 11 (64%) non-domestic felids that originally tested FeLV-positive,
were negative when retested. The remaining four were not retested and did not go on to
develop clinical signs of FeLV. Clinical signs in non-domestic felids with self-limiting
infections were minimal and included lethargy, peripheral lymphadenopathy, and
dehydration. Terminal infections were seen in a free-ranging and captive puma, a bobcat,
and a cheetah. Clinical pathology and necropsy findings included anemia, lymphopenia,
other cytopenias, septicemia, lymphadenopathy, opportunistic infections, and lymphoma
(Meric, 1984; Jessup et al., 1993; Sleeman et al., 2001; Marker et al., 2003).
In Florida panthers, routine FeLV ELISA antigen testing was negative since testing
began in 1978 through late 2002 (Roelke et al., 1993b; Florida Fish and Wildlife
Conservation Commission, unpubl. data); however, during the 2002-2003 capture season,
two panthers tested antigen-positive. These findings launched the investigation detailed
in this report.
The objectives of this study were to determine for FeLV in Florida panthers 1) the
history of exposure, 2) the prevalence and geographic distribution, 3) the outcome
following exposure, 4) the clinical signs, clinical pathology, and pathological changes
associated with infection, and 5) risk factors for infection.
-65% eventually clear virus
Rovir + to+++
Antigen+ 3-13 wks
Rovir -++ to ++
IFA + Life
~35% of those exposed will
remain persistently infected
Adaptedfrom: Hartmann(2005), Hoover
and Mullins (1991), and Torres et al. (2005).
Figure 1-1. Outcome following exposure to feline leukemia virus in domestic cats.
MATERIALS AND METHODS
Study Area and Period
Florida panthers were sampled in southern peninsular Florida (south of 280 N)
primarily in the Big Cypress and Everglades ecosystems. For ELISA antibody
comparisons, capture/sampling locations were divided into north and south of 1-75
(approximately 28.050 N) (Fig. 2-1). The prospective study period was 1 July 2002 to 5
June 2005. Archived tissues collected between 1990 and 30 June 2002 were
retrospectively evaluated. For analysis of ELISA antibody prevalence, the study period
was divided into before (1990-1995) and after (1996-2005) genetic restoration.
Previously published and unpublished FeLV ELISA antigen test results from 1983 to 30
June 2002 are included in this report (Roelke, 1990; Roelke et al., 1993b; Dunbar, 1994;
FWC, unpubl. data).
Florida Panther Capture and Immobilization
Free-ranging Florida panthers and translocated Texas cougars were captured using
trained hounds. Panthers either bayed on the ground or were treed, and then were darted
with a 3 ml compressed-air dart fired from a C02-powered rifle. Since 2002,
immobilization drugs included various combinations of ketamine HC1 (Congaree
Veterinary Pharmacy, Cayce, South Carolina, USA), medetomidine (Domitor, Pfizer
Animal Health, Exton, Pennsylvania, USA), tiletamine HCl/zolazepam HC1 (Telazol,
Fort Dodge Animal Health [FDAH], Fort Dodge, Iowa, USA), midazolam HC1 (Abbott
Laboratories, North Chicago, Illinois, USA), and xylazine HC1 (Congaree Veterinary
Pharmacy, USA) (Shindle et al., 2003; Shindle et al., 2004). Following immobilization,
treed panthers were caught with a net and, in some cases, a crash bag (McCown et al.,
1990). Propofol (PropoFloTM, Abbott Laboratories, North Chicago, Illinois, USA) was
administered intravenously (IV) either as a bolus or continuous drip to maintain
anesthesia. Butorphanol tartrate (0.1-0.4 mg/kg, FDAH) or midazolam HC1 (0.03 mg/kg)
was administered intramuscularly (IM) to smooth recovery in some panthers. Panthers
were left to recover in a shaded area away from water. Xylazine HC1 and medetomidine
HC1 were reversed with yohimbine HC1 (Yobine, Lloyd, Inc., Shenandoah, Iowa, USA)
and atipamezol HC1 (Antisedan, Pfizer Animal Health, Exton, Pennsylvania, USA),
respectively, at 12 to 14 their recommended dosages.
Vital signs (temperature, heart rate, respiration rate, and capillary refill time) and
depth of anesthesia were monitored and recorded. A sterile petrolatum ophthalmic
ointment was applied to the eyes for lubrication. All animals underwent a physical
examination to assess general health and physical condition. For each panther handled,
the skin over the medial saphenous vein was clipped, prepped, and an IV catheter
aseptically placed. Sterile isotonic fluids were administered either subcutaneously (SQ)
or IV. Panthers were implanted with a SQ transponder identification chip (Trovan,
Douglas, United Kingdom), ear-tattooed, measured, and weighed.
Live-capture Sample Collection
Approximately 70-140 ml of blood (depending on body weight) were collected
from the medial saphenous or cephalic veins using a butterfly catheter (19 or 21 gauge),
luer adapter/hub, and Vacutainer tubes (Becton Dickinson, Franklin Lakes, New Jersey,
USA) (approximately 50 ml in serum separator, 40 ml in EDTA, 9 ml in Na Heparin, and
9 ml in ACD tubes). From uncollared panthers, eight skin biopsies (4 mm) were collected
aseptically from the medial aspect of the hindlimbs and saved in biopsy transport media.
Defects were closed with surgical glue. Hair clipped from blood collection and biopsy
sites and pulled hair were saved in sample collection bags; clipped hair was saved also
from the ventral abdomen. Other samples such as bacterial cultures, skin scrapings, and
diagnostic biopsies were taken if indicated. Between November 2002 and April 2004,
blood smears were made from EDTA whole blood on glass slides approximately 6 to 24
hr after collection. Beginning May 2004, blood smears were made in the field from fresh
Vaccination and Treatment
Panthers >4 mo old were vaccinated SQ against feline viral rhinotracheitis (FVR),
feline calicivirus (FCV), feline panleukopenia virus (FPV) (Fel-O-Vax PCT [FDAH], 1
ml, lower left leg), and rabies (RabvacTM 3 [FDAH], 1 ml, lower right leg). Beginning
June 2003, captive and free-ranging panthers were also vaccinated against feline
leukemia virus (FeLV, Fel-O-Vax Lv-K [FDAH] or Fevaxyn FeLV, Schering-Plough
Animal Health Corporation, Omaha, Nebraska, USA, 2 ml, lower left leg). Some
panthers were given a FeLV booster (2 ml) IM remotely by darting 3-16 wk post initial
inoculation. Captured panthers were dewormed with ivermectin (0.1 mg/kg, Ivomec,
Merial Limited, Iselin, New Jersey, USA) and praziquantel (3.75 mg/kg, CestaJectTM
Phoenix Pharmaceutical, Inc., St. Joseph, Missouri, USA) administered SQ in the lateral
aspect of thigh. Penicillin G procaine/benzathine (USVet, Hanford Pharmaceuticals,
Syracuse, New York, USA) was administered IM at 22,000 to 44,000 U/kg.
Captured adult and juvenile panthers were fitted with a VHF or VHF/GPS radio-
collar and monitored three times weekly as described by Shindle et al. (2004). If a
mortality signal was detected the carcass was recovered the same day for necropsy.
Neonatal kittens were handled according to Land et al. (1998) and marked with a
SQ transponder identification chip. Pyrantel pamoate (22 mg/kg, Anthelban V, Phoenix
Pharmaceutical, Inc., St. Joseph, Missouri, USA) was administered orally. Blood was
collected from the jugular vein.
All FeLV-positive Florida panthers and/or those found dead due to infectious
disease or unknown causes were completely necropsied by board-certified pathologists at
the University of Florida (Veterinary Medical Teaching Hospital, Gainesville, Florida,
USA) or Disney's Animal Kingdom (Celebration, Florida, USA). One severely autolyzed
FeLV-positive panther (FP109) and all panthers dying of known trauma were necropsied
by the FWC veterinarian at the Wildlife Research Laboratory (FWC, Gainesville, Florida,
When carcass condition allowed, tissue samples were collected at necropsy from all
major organs. Fluids collected included heart blood, venous blood, thoracic blood,
aqueous humor, and urine. Blood samples were centrifuged at 2000 rpm for 10 minutes
and the supernatant decanted. Representative tissues from fresh (unfrozen) and some
previously frozen panthers were placed in 10% neutral buffered formalin. Fixed tissues
were embedded in paraffin, sectioned at 5 to 6 [tm and stained with hematoxylin and
All tissues from live-captured and necropsied panthers not immediately analyzed
were archived at -200 to -700C.
Age Determination and Genetic Status
Panther ages were either known (handled as kittens) or were estimated from tooth
wear. Panthers were classified as neonates (<8 wk-old), dependents (8 wk to <1 yr),
subadults (1 to <2.5 yr), adults (2.5 to <10 yr), and older adults (>10 yr).
Panthers were grouped by genotype (canonical Florida panther, Texas puma, Texas
puma/Florida panther intergrade, Texas puma/Everglades/Florida panther intergrade,
SIR/Florida panther intergrade, and Everglades/Florida panther intergrade) (W. Johnson,
Enzyme-linked Immunosorbent Assay Antibody
Antibodies to FeLV were detected at Hansen Veterinary Immunology (Dixon,
California, USA) according to techniques described by Lutz et al. (1980b). Optical
densities (OD) of less than 0.25 were considered negative, 0.25 to <0.35 were low
positive, 0.35 to <0.5 were medium positive, and those >0.500 were high positive. For
statistical analysis any OD >0.25 was considered positive.
Enzyme-linked Immunosorbent Assay Antigen
Serum for ELISA antigen testing (ViraCHEK FeLV, Synbiotics Animal Health,
San Diego, California, USA) was shipped overnight to the New York State Diagnostic
Laboratory (Comell University, Ithaca, New York, USA). Adsorbing reagents were used
to remove heterophile antibody. Fluids collected from live-captured and necropsied
panthers were tested for FeLV antigen with a rapid immunochromatic assay (SNAP
Combo, IDEXX Laboratories, Westbrook, Maine, USA). Beginning November 2003,
EDTA whole blood from captured panthers was tested in the field using the SNAP
combo. The SNAP Combo was also used to test fluids collected from necropsied
panthers. Fluids included blood collected from the thoracic cavity, heart chambers,
vessels, and marrow cavity, and aqueous humor.
Immunofluorescent Assay and Immunohistochemistry
Panthers testing positive by ELISA antigen were also tested by IFA.
Immunofluorescent assays were performed on EDTA or fresh whole blood smears using
techniques described by Hardy et al. (1973) at the National Veterinary Laboratory
(Franklin Lakes, New Jersey, USA). Immunohistochemistry to identify p27 antigen was
performed on formalin-fixed paraffin-embedded tissues at the Diagnostic Center for
Population and Animal Health (Michigan State University, Lansing, Michigan, USA)
using a labeled streptavidin-biotin peroxidase detection system on an automated stainer
(Ramos-Vara et al., 2002).
Polymerase Chain Reaction, Genetic Sequencing, and Viral Culture
Polymerase chain reaction and subsequent genetic sequencing was performed at the
Laboratory for Genomic Diversity (National Cancer Institute, Frederick, Maryland, USA)
on tissues collected from panthers at capture and necropsy. Viral culture was performed
at the Center for Retrovirus Research (The Ohio State University, Columbus, Ohio,
USA). Materials and methods used, and complete results for PCR and genetic sequencing
(M. Brown, unpubl. data) and viral culture (K. Hayes, unpubl. data) will be presented in
Complete Blood Count and Serum Chemistry
Complete blood counts (CBC) and serum biochemical parameters were determined
by Antech Diagnostics (Smyrna, Georgia, USA). Blood smears were examined at the
Veterinary Medical Teaching Hospital (University of Florida, College of Veterinary
Medicine, Gainesville, Florida, USA) for hemoparasites, white blood cell differential
counts, and red blood cell morphology.
Other Diagnostic Testing
Necropsied panthers were tested for rabies by IFA at the Jacksonville Central
Laboratory (Jacksonville, Florida, USA). Viral isolation and real-time and conventional
PCR for canine distemper virus (CDV), pseudorabies virus, flaviviruses, and alphaviruses
were performed at the Southeastern Cooperative Wildlife Disease Study (Athens,
Georgia, USA) on brain, heart, and other tissues collected from panthers dying of
Other serological tests included Western blot for FIV and kinetics-based enzyme-
linked immunosorbent assay (KELA) for feline coronavirus antibodies (FCV) (New York
State Diagnostic Laboratory). Polymerase chain reaction for Mycoplasma haemofelis and
M. haemominutum was performed on EDTA whole blood from FeLV positive panthers
at the University of Illinois (College of Veterinary Medicine, Urbana, Illinois, USA) and
Cornell University (Ithaca, New York, USA).
Prevalence was calculated as the percentage of panthers/pumas positive for FeLV
antibodies by ELISA (OD >0.251). Raw prevalence estimates were examined for each
potential categorical predictor (age classes, genotype, FIV status, location, time period,
and gender). Logistic regression using Egret software (Cytel Software Corporation,
Cambridge, Massachusetts, USA) was performed to investigate ELISA antibody status as
a binary response variable. Odd ratios and their 95% confidence limits were calculated
for each state of the categorical predictors in comparison to an arbitrary reference state.
Significance of difference from 1.0 was determined for the odd ratios by the Wald test.
To account for correlation among replicate outcomes from individuals with multiple test
results, panther identification was modeled as a random effect within the logistic
regression model. Significance of the random effect was evaluated by a likelihood ratio
test. Test results were considered significant at P<0.05. The two significant predictors
emerging from univariate analyses (location and time period) were included in a multiple
predictor logistic regression analysis and their interactions examined.
SF r, .. i:
* Public lands
F[ Private lands
Figure 2-1. Study area in south Florida, USA.
Enzyme-linked Immunosorbent Assay Antibody
ELISA antibody ODs were determined for samples collected from 128 Florida
panthers/Texas pumas on 257 occasions between 2 January 1990 and 29 March 2005.
Eighteen (7%) samples from 17 individuals were positive (1 high OD, 3 medium OD, 14
The prevalence of positive antibody ODs was significantly greater in the period
1996-2005 compared to 1990-1995 (P = 0.032). The prevalence of positive antibody ODs
was significantly greater among panthers sampled north of 1-75 compared to south (P =
0.014). No positive ODs were found in the southern portion of panther range (south of
US41). The odds of having a positive antibody OD were not affected by age, gender,
genotype, or FIV status. Of panthers sampled on multiple occasions, six had low or
medium positive ODs at their initial sampling but seroconverted to negative status when
re-sampled 10 mo to 3 yr later.
0' 1J J W Li North
AWam, CZ LJ. NC
90 91 92 93 94 95 9697 4w Aw 4 W Central
96 9 98 99 O0 I T South
02 03 04 05
Figure 3-1. Distribution of positive feline leukemia virus positive ELISA antibody optical
densities in Florida panthers/Texas pumas by region and year 1990-2005. North
refers to lands north of CR846, NC refers to lands between CR846 and 1-75,
Central refers to lands between 1-75 and US41, and South refers to lands south of
Enzyme-linked Immunosorbent Assay Antigen
Prior to the study period, all (n = 143 sampled on 322 occasions) Florida panthers
and Texas pumas sampled were negative for p27 antigen by ELISA based on review of
published and unpublished data and retrospective testing. During the study period (1 July
2002 to 5 June 2005), 91 panthers/Texas pumas were tested on 113 occasions for FeLV
antigen by ELISA. Fifty-five panthers or pumas were sampled on 66 occasions at
capture, 40 were sampled at necropsy, 10 were sampled at both capture and necropsy,
and seven were tested as neonatal kittens. Panther number, age, gender, FIV status, and
results of FeLV diagnostic tests are presented in Table A-1.
The prevalence of antigenemia (positive ELISA antigen) among panthers and
Texas pumas >1 yr of age, not previously vaccinated for FeLV, and sampled during the
study period, was 7% (5 of 71). All antigenemic panthers were captured in
Okaloacoochee Slough (OKS). The prevalence of antigenemia in OKS (NC region, Fig.
3-2) was 45.5% (5 of 11).
Antigenemia was only detected in adult panthers (3 males, 2 females). The average
age of antigenemic panthers was 4.85 yr (standard deviation [SD]+3.5) and ranged from
2.25 to 11 yr. Genotypes included canonical Florida panthers (n = 3), Texas puma/Florida
panther intergrade (n = 1), and SIR captive/Florida panther intergrade (n = 1). Case
histories of antigenemic panthers are presented in Appendix B.
Feline leukemia virus antigen was detected by SNAP test in all fluids tested in
those viremic panthers suitable for testing at necropsy. Fluids testing positive included
thoracic blood (FP115, 122, 123, 132), splenic blood (FP115), venous blood (FP132), and
aqueous humor (FP115, 122, 123, 132).
Immunofluorescent Assay and Immunohistochemistry
Three (FP122, 123, 132) of the 5 (60%) panthers positive for FeLV antigen by
ELISA were also IFA positive. Results for two viremic panthers (FP109, 115) were
inconclusive. Spleen and lymph node from 2 of 2 (100%) viremic panthers (FP115, 132)
were positive for p27 antigen by IHC.
During the study period, 37.5% of panthers/Texas pumas tested were positive for
FIV antibodies by Western blot. Three of five (60%) FeLV antigen-positive panthers also
tested positive for the puma lentivirus strain of FIV (J. Troyer, unpubl. data). Serology
for FCV was negative for all panthers sampled during the study period (n = 64).
Clinical signs observed at capture in four antigenemic panthers included a
peripheral lymphadenopathy (n = 2, 50%) and muscle wasting (n = 1, 25%).
Complete blood counts were performed on four antigenemic panthers sampled
while living. Significant findings included a mild to moderate non-regenerative anemia (n
= 3 [75%]), lymphopenia (n = 3), low hemoglobin (n = 3), monocytosis (n = 1 [25%]),
and elevated nucleated red blood cell count (n = 1). Large immature mononuclear cells
with prominent nucleoli, consistent with acute lymphoblastic leukemia, were seen in two
panthers (FP122, 123, 50%). The mean hematocrit of antigenemic panthers was 29.3%
(SD+ 7.9, range 22.5-42.5%), hemoglobin 9.3 g/dL (SD2.4, range 7.2-13.2 g/dL), red
blood cell count 6.2 x 106/tl (SD+1.78 x 106/tL, range 4.2-8.75 x 106/tl), and
lymphocyte count 1165/4l (SD811.9/4l, range 490-2250/4l). Serum biochemical values
in antigenemic panthers were unremarkable. Clinical pathology of antigenemic panthers
and normal values for panthers are summarized in Table B-1.
Three antigenemic panthers (FP115, 122, 132) were suitable for complete necropsy
based on carcass condition. Completely necropsied panthers had evidence of anemia
(pale mucus membranes and skeletal muscle, n = 2 [66.7%]), moderate to severe
dehydration (n = 2), lymphadenopathy (n = 2), septicemia (n = 2), bronchointerstitial
pneumonia (n = 2), abscesses (n = 1, [33.3%]), and puncture wounds (n = 1). Lacerations
and puncture wounds associated with intraspecific aggression (ISA) were seen in the two
autolyzed/decomposed carcasses (FP109, 123).
Microscopic examination was performed on three panthers (FP115, 122, 132).
Sections of bone marrow from two (66.7%) panthers were hypercellular with
approximately 90 to 100% of the marrow space occupied by hematopoietic cells.
Megakaryocytes were present in normal to moderately increased numbers. No marrow
evidence of acute lymphoblastic leukemia was seen. Microscopic changes consistent with
septicemia were seen in most tissues in FP115 and 132. See Appendix B for complete
Aerobic culture of multiple tissues from FP115 and 132 resulted in heavy growth of
Escherichia coli and P-hemolytic Streptococcus sp., respectively. Rabies IFA and viral
isolation and PCR for canine distemper virus, pseudorabies virus, flaviviruses, and
alphaviruses were negative in FP122.
Two of four (50%) ELISA antigen-positive panthers (FP109, 115) were PCR
positive for M. hemominutum; FP115 was also positive for M. haemofelis (J. Messick,
unpubl. data). Organisms were not seen on blood smears made from EDTA whole blood.
Rare Cytauxzoon felis organisms were seen on blood smears from FP109; blood smears
from FP115, 122, and 123 were negative.
Suspected causes of death for the five antigenemic panthers included septicemia (n
= 2), intraspecific aggression (n = 2), and anemia/dehydration (n = 1). Time from
diagnosis to death averaged 9.25 (SD+10.3, range 2-24.6) wk in panthers antigenemic at
capture (FP109, 115, 122, 123). Time from diagnosis to death in the two panthers
believed to have died due to FeLV-related diseases was 2 (FP122) and 24.6 wk (FP115).
Time from exposure to death for one panther (FP132) dying of FeLV-related disease was
Figure 3-2. Feline leukemia virus (FeLV) ELISA antigen results for panthers >1 yr, not
previously FeLV vaccinated, and sampled in South Forida (south of
Caloosahatchee River) between 1 July 2002 and 5 June 2005.
Diagnostic tests validated for domestic animals but used on wildlife must be
interpreted with caution (Hietala and Gardner, 1999). Nevertheless, the test results in this
study were biologically consistent and appeared to be appropriate and suitable for use in
The ELISA antibody test detects exposure to FeLV and is considered more
sensitive but less specific than Western blot analysis. ELISA antibody testing has only
been used rarely in non-domestic felids. Ryser-Degiorgis et al. (2005) found serum from
58 of 102 (58%) Eurasian lynx (L. lynx) to be FeLV positive by ELISA antibody but
negative by Western blot. The authors speculated that cross-reactions with E. coli antigen
(test preparation) or antibodies to murine leukemia viruses may have been responsible for
the false-positive results. Our positive ELISA antibody tests were not confirmed by
Western blot. Further, antibody ODs in panther serum were tested incrementally.
Incremental testing of serum for ELISA antibodies may lead to inconsistent results due to
between-batch variation. Additionally, degradation of antibodies in stored serum may
have resulted in the apparent increase in positive ODs in recently collected samples.
However, FeLV antibodies are stable when frozen in serum (S. Hansen, pers. commun.).
Additionally, ELISA antibody results in Florida panthers were consistent biologically
with other test results and observations. Panthers seroconverted following vaccination
(data not shown) and positive ODs were geographically and temporally clustered.
ELISA antigen tests detect the FeLV p27 protein and therefore should be suitable
for use in exotic species. Nevertheless, false-positives have occurred in some tests that
used murine-derived reagents in domestic and non-domestic cats that had naturally
occurring anti-murine antibodies (Lopez and Jacobson, 1989). False positive results were
reported in one Florida panther tested in 1987 (Lopez, 1988). In this case anti-mouse
antibodies were believed to have resulted from vaccination with a rabies vaccine of
mouse brain origin. Changes in test procedures and reagents effectively eliminated this
problem by the early 1990s (Jacobson and Lopez, 1991). False positives may occur also
due to insufficient washing of vessels in micro-well systems (Jarrett et al., 1982), a
problem not encountered when using rapid immunoassay test kits.
The effectiveness of using body fluids from known infected panthers for detection
of p27 antigen was evaluated. Hemolyzed thoracic, heart, and venous blood; bone
marrow; and aqueous humor from infected panthers consistently tested positive by rapid
immunoassay (SNAP Combo), even on severely autolyzed specimens. The p27 antigen is
only 27,000 daltons and is thus small enough to cross into the aqueous humor in healthy
felids (K. Gellatt, pers. commun.). Thus aqueous humor, and the other fluids described
above, may be useful for FeLV monitoring not only in panthers but in other populations
Immunofluourescent assay and IHC detect p27 antigen in platelets and neutrophils
of blood smears and paraffin-embedded fixed tissues respectively. Three of 5 panthers
positive by ELISA antigen were also positive by IFA; two (FP109, 115) were
inconclusive. Inconclusive results in these panthers may have been due to a delay in
testing and/or improper slide storage. Alternatively, if the samples were true negatives,
sampling may have occurred soon after exposure, before infection of the bone marrow. In
domestic cats neutropenia or thrombocytopenia can lead to false negatives; however,
these values were normal in FP109 and FP115. Spleen and lymph node from FP115 did
test positive by IHC when collected 5 mo after initial positive antigen findings. However,
tissues from FP109 were severely autolyzed and unsuitable for IHC when collected at
necropsy approximately 1 mo after initial positive antigen findings.
History of Exposure
The FeLV epizootic in free-ranging Florida panthers was foreshadowed by
evidence of increasing exposure based on ELISA antibody tests. Beginning in the late
1990s the prevalence of positive ODs increased dramatically, peaking in 2001 when 9 of
26 (34.6%) were positive (Fig. 3-1). Positive ODs were also geographically clustered
with 16 of 18 (88.9%) positive ODs occurring north of 1-75 (Fig. 3-1). In domestic cats,
antibody ODs increase with age; however, this was not seen in panthers probably the
result of small sample size.
ELISA antibody tests support the theory of multiple introductions of the virus. One
introduction may have occurred on the Florida Panther National Wildlife Refuge
(FPNWR) between January and November of 2001. Four of five (80%) panthers sampled
during the spring of 2001 (2000-2001 capture season) were negative for ELISA
antibodies. Two of these panthers (FP96, 99) were recaptured the next season (2001-
2002). FP96 had seroconverted from a negative to low positive OD, and FP99 went from
a low to medium positive OD. Two other panthers captured in the Fall of 2001 also had
positive ODs (FP107, low; FP78, medium). Based on telemetry data, FP96 and 107
(siblings) and FP99 formed a loosely associated group between August and December
2001 (Land et al., 2002). This may have facilitated exposure among these panthers if any
were shedding virus at the time. Indeed, one of these, FP96, was found to be latently
infected (PCR positive, M. Brown, unpubl. data) at necropsy after being killed by another
male in early 2002. Although no panthers from FPNWR tested antigen-positive when
sampled, we speculate that FP96 became transiently viremic and exposed the panthers
that were accompanying him. He apparently overcame the infection, perhaps aided by a
relatively high antibody OD.
Prevalence and Distribution
Before the 2002-2003 capture season, routine ELISA antigen testing of captured or
necropsied Florida panthers had been negative since 1978. However, between July 2002,
and June 2005, 5 of 71 (8%) free-ranging panthers/pumas >1 yr of age sampled had
active FeLV infections based on ELISA antigen, IFA, and/or viral culture results. All
infected panthers had overlapping home ranges in the OKS ecosystem in the north-central
portion of panther range (Fig. 3-2).
Outcome Following Exposure
In domestic cats, prolonged exposure is generally necessary for transmission.
Indeed the percentage of adult domestic cats becoming persistently infected following a
single exposure event is only 3% (Hartmann, 2005). However, FP132 became infected
apparently after an aggressive encounter with an infected panther (FP123). At
examination approximately 2 days after the fight, FP132 had only minor scratches and
two puncture (presumably bite) wounds. Although FP132 was FeLV antigen negative at
this time, he developed a persistent FeLV infection and died 4 mo later. We speculate
that bite wounds are an important mode of FeLV transmission in panthers. Feline
leukemia virus is present in highest concentrations (106 infectious units/ml) in the saliva
(Francis et al., 1977). A relatively larger dose of saliva would be expected to be
transferred between fighting panthers versus domestic cats and may explain the apparent
ease of transmission.
The presence of infection in females is evidence that transmission also may occur
during breeding. Males and females will pair for 2 to 5 days and transmission may occur
during copulation, mutual grooming, or biting. Although FeLV virus is present in the
semen of domestic cats, venereal transmission is not considered important (Hoover and
In many respects, the outcome following exposure to the virus in panthers is similar
to that in domestic cats. Following exposure, a panther can clear the virus early
(abortive/transient infections) or can become latently or persistently infected.
Based on the relatively large number of panthers with positive ELISA antibody
ODs but antigen and PCR negative (M. Brown, unpubl. data) test results, many panthers
exposed to the virus are able to clear the infection soon after exposure. Assuming a
similar pathogenesis to that occurring in domestic cats, panthers in this category would
have cleared the infection within several weeks of exposure before infection of the bone
marrow. The majority of domestic cats in this category are considered refractory to re-
infection (Hardy, 1980b). Based on telemetry data, at least one female (FP110) with
evidence of a previous abortive/transient infection survived exposure to at least two
FeLV positive males without developing persistent viremia.
It is possible that FP109 was transiently infected when captured in January 2003.
At capture he was anemic, lymphopenic, and had a profound lymphadenopathy. Levy
(1999) described similar signs in transiently infected domestic cats. FP109 also had a
high ELISA antibody OD. Antibodies detectable by ELISA appear shortly after infection
in domestic cats (Lutz et al., 1980b), and high antibody ODs in domestic cats are a good
prognostic indicator for recovery (Hofmann-Lehmann et al., 2001).
There is also evidence that some panthers can become latently infected as
evidenced by positive PCR and antibody ODs but negative ELISA antigen tests. These
panthers presumably failed to control infection until later in the course of infection, and
retained provirus in leukocytes sufficient to be detectable by PCR. No latently infected
panther has developed a persistent infection, and at least one latently infected panther is
still surviving in the wild at least 2 yr after diagnosis.
In domestic cats, persistent infection is usually characterized by bone marrow
infection (positive IFA), viremia persisting for >16 wk, and eventual FeLV-related
clinical signs. A diagnosis of persistent infection in panthers was also based on test
results, duration of infection, and clinical signs; however, premature deaths, severe
autolysis, and limited ability to re-sample panthers while living precluded complete
determination of disease progression. Nevertheless, persistent infections were diagnosed
in four panthers (FP115, 122, 123, and 132). These diagnoses were based on viremia >16
wk (FP 115), positive IFA (FP122, 123, 132), and the presence of FeLV-related diseases
(FP115, 122, 123). The latter criterion is speculative; FeLV-infected domestic cats are
subject to the same diseases as non-infected cats (Levy, 1999). Nevertheless, the finding
of septicemia in two necropsied infected panthers and apparent acute lymphoblastic
leukemia in two live-captured infected panthers appears to be unique to those infected
with FeLV. Septicemia without apparent cause or neoplasia was not observed in 73
panther necropsies performed 1978-1999 (Taylor et al., 2002).
Persistently infected panthers had relatively low antibody ODs. Although FP115
had a medium positive OD, FP122, 123, and 132 had negative ODs suggesting a muted
humoral response to infection. In domestic cats, low ELISA antibody ODs are
characteristic of persistent infections (Hoffmann-Lehmann, 2001).
Immunosuppression is a common feature of FeLV infection and is believed to
result in increased susceptibility to infectious diseases. Co-infections were the most
frequent finding in FeLV infected domestic cats examined at North American veterinary
schools (Levy, 1999). Infectious and parasitic diseases seen more commonly in FeLV-
infected domestic cats than non-infected cats included bacterial infections,
hemobartonellosis (Mycoplasma spp.), FCV, upper respiratory infections, babesiosis,
stomatitis, coccidiosis, and toxoplasmosis (Grant et al., 1980; Reinacher, 1989; Reinacher
et al., 1995). Of these, bacterial infections, including p-hemolytic streptococci, were most
important. Jessup et al. (1993) diagnosed septicemia and leptospirosis in a FeLV-infected
puma from California.
The most significant apparent opportunistic infections in panthers were bacterial;
FP 115 had an E. coli septicemia while a mixture of opportunistic bacteria, predominately
0-hemolytic streptococci, was cultured from FP132. Other opportunistic infections in
viremic panthers may have included M. haemofelis and M. haemominutum; however,
approximately 70% of FeLV-negative Florida panthers also tested positive for these
mycoplasmas (J. Messick, unpubl. data). Feline infectious peritonitis has not been
diagnosed in panthers regardless of FeLV status.
Anemias, primarily non-regenerative, are also a frequent finding in FeLV-infected
domestic cats. Non-regenerative anemias were seen in FP109 and FP122 when live
captured, and may have been the cause of death in FP122. Severe anemia in FP132 was
also suspected at necropsy. Bone marrow sections from FP122 and FP132 were examined
histologically. Sections were hypercellular with approximately 90% of the marrow space
occupied by hematopoietic cells; however, erythroid precursors were decreased in
number, and few maturing erythroid cells were present.
Finally, hematopoietic neoplasias occur frequently in domestic cats (Reinacher,
1989). Under controlled conditions opportunistic infections can be reduced and neoplasia
becomes the most important cause of mortality in FeLV-infected cats (Hofmann-
Lehmann et al., 1997). Given the apparently rapid progression of infection, FeLV-
positive panthers may not have survived long enough to develop terminal neoplasia.
Atypical lymphocytes consistent with acute lymphoblastic leukemia were seen on blood
smears from FP122 and FP123; however, no evidence of leukemia was seen on
histological examination of bone marrow or other organ tissue from FP122 (FP123 was
unsuitable for histological examination). The role, if any, in the deaths of these panthers
All antigenemic panthers died relatively soon after diagnosis. FP115 and FP132
died from septicemias (E. coli and P-hemolytic streptococci respectively). FP122 is
believed to have died from severe anemia. FP109 and FP123 died from ISA; however,
anemia or other FeLV-related diseases (acute lymphoblastic leukemia) may have
impaired their ability to fight. For example, FP109 had a hematocrit of 24% (normal is
36.4% [Dunbar et al., 1997]) when handled 1 mo prior to death, which may have resulted
in exercise intolerance. Although FP123 was apparently healthy when handled 6 wk prior
to death, he inflicted only a few minor punctures and scratches to the panther that killed
him (FP132). Further, due to severe autolysis, it is unknown if these panthers may have
died from secondary bacterial infections. Neither FP109 nor FP123 had obviously fatal
ISA-related wounds. Of concern was the presence of healing wounds associated with ISA
in FP132 at the time of death suggesting that he may have exposed another panther
before dying. Further, these bite wounds may have been the source of infection leading to
septicemia in this panther.
Progression of infection appears to be quite rapid in panthers. Although 50% of
viremic domestic cats die within 6 mo of exposure (Jarrett, 1983), adult cats enjoy a
longer induction period and less severe disease compared to younger age groups (Hoover
et al., 1976; Levy, 1999). All viremic panthers were adults and, although the time of
infection is unknown in most infected panthers, the average time from diagnosis to
mortality was just over 9 wk. In the one case of known exposure, FP132 died 18 wk after
exposure to an infected male. Lack of supportive care and presumably increased exposure
to pathogens may play a role in this apparently more rapid clinical course.
Progression to persistent infection following exposure depends on a number of host
and viral factors. The most important host factor in domestic cats is age, but inbreeding
(genetic variation) and FIV status also affect the outcome. Important viral factors include
strain, dose, and duration of exposure. However, with the average age of viremic panthers
approaching 5 yr, maturity did not appear to be protective against infection. Genetic
variation also did not appear to significantly influence the outcome following exposure.
While some panthers had very low genetic variation, at least two had He much greater
than the average for the population (M. Roelke, unpubl. data). Ancestry also did not
appear to play a role. Although three viremic panthers had canonical or original
genotypes, two were intergrades (Florida panther/Texas puma and Florida panther/SIR
captive) (W. Johnson, unpubl. data).
Feline immunodeficiency virus and FeLV, both retroviruses, have overlapping host
cell tropism. In domestic cats, co-infection with FIV results in marked synergism of
immunosuppression and clinical disease induction (Pedersen et al., 1990). Three of five
(60%) FeLV viremic panthers were co-infected with FIV (Shindle et al., 2003; J. Troyer,
unpubl. data); however, the impact of co-infection in the panther is unknown.
Historically, approximately 28% of the free-ranging panther population was
infected with the puma-lentivirus strain of FIV (Olmstead et al., 1992). However, there is
a disturbing trend in FIV prevalence 76% (13 of 17) of panthers captured during the
2004-2005 capture season were positive for FIV antibodies. The consequences of this
trend are unknown but if pre-existing FIV infection affects subsequent FeLV infection,
then an increasing FIV prevalence may alter the epizootiology of FeLV infection in
Viral factors may play a more important role in the apparent greater impact of
FeLV on panthers. Although infection pressure is expected to be low in this reclusive
solitary species, viral load may be quite high. Although the virus concentration in saliva
of infected panthers is unknown, the dose transmitted during fighting or breeding would
be expected to be higher as panthers may be as much as 15-fold larger than domestic cats.
However, virus strain may be the most important factor in the current epizootic. Based on
preliminary viral culture results, the strain isolated from the current epizootic in panthers
may be similar to virulent domestic cat strains (K. Hayes, unpubl. data).
The source of infection in panthers is unknown. Texas pumas introduced in 1995
tested antigen negative during quarantine (Dunbar, 1995), and western pumas in captivity
at SIR tested negative when handled in 1999 (Shindle et al., 2000). In reports of FeLV
infection in non-domestic felids, authors speculated or provided direct evidence that an
infected domestic cat was the source and this is the most likely explanation for FeLV in
panthers. Domestic cat remains have been found in the stomachs of necropsied pumas
from California (Jessup et al., 1993) and there have been observations of panthers killing
domestic cats in Florida (L. Richardson, pers. commun.). Kennedy-Stoskopf (1999)
speculated that consumptionin of FeLV infected domestic cats by larger nondomestic
felids would...be an effective way to transmit the virus."
Private land in panther range continues to be developed at an astounding pace. As
humans encroach on panther habitat they are accompanied by their domestic animals,
including cats. Indeed free-roaming domestic cats have been observed on private lands
near OKS (M. Lotz, pers. commun.). Additionally, the increasing panther population
undoubtedly results in increased opportunities for exposure to domestic cats. Young
dispersing males move through the fringes of the resident population and often occupy
marginal habitat until an established home range becomes available (Maehr, 1997). This
existence on the urban/wildland interface likely increases the risk for exposure to
domestic cats. Riley et al. (2004) speculated that a higher prevalence of positive feline
calicivirus titers in bobcats frequenting urban areas was due to increased exposure to
The transmission of FeLV from a domestic cat to a panther is likely a rare event.
Domestic cat remains have never been reported in panther scat (Maehr et al., 1990) or in
stomach contents (M. Cunningham, unpubl. data). Further, the odds of a free-ranging
domestic cat in Florida having FeLV is less than 1 in 20 (Lee et al., 2002). Should these
unlikely events occur the panther would still need to become persistently infected
Given that these events, however unlikely, probably did occur, once the species
barrier was crossed the virus was likely spread panther-to-panther. The apparent
transmission of FeLV from FP123 to FP132 supports this theory. Higher panther
densities undoubtedly facilitate this panther-to-panther transmission. The population has
tripled since the early 1990s while panther habitat has been reduced.
Feline leukemia virus infection in panthers is likely a disease of adult cats.
Although mother-to-offspring transmission is probably the most important mode of
transmission in domestic cats (Levy, 2005) it is unlikely to be a factor in the
epizootiology of the disease in panthers. Given the apparent severity and rapid
progression of the disease in panthers, infected females are unlikely to survive to raise
kittens. Further, if a female were to successfully reproduce, infected kittens would be
unlikely to survive to independence the age at which an infected kitten would first be
expected to encounter susceptible panthers and potentially spread the virus.
The current epizootic likely began in the OKS area in late 2001 or early 2002,
possibly as a result of cross-species transmission from an infected domestic cat on a local
ranch. The first positive antibody tests in OKS began to appear in February 2002. The
infection was then likely spread panther-to-panther resulting in the infection of at least
five panthers. However, since July 2004, none of 30 panthers have tested FeLV antigen-
positive indicating that the epizootic may be over. Several factors may have contributed
to this. First, the rapid progression of disease may have limited the number of exposure
events infected panthers die before transmitting the disease. Additionally, some
panthers appear to be refractory to infection thus limiting the number of susceptible
individuals capable of perpetuating the disease. Small population size and geography may
have also helped. Fromont et al. (1998a) demonstrated that populations less than 100
individuals were unlikely to sustain FeLV infections. Since this epizootic occurred in the
northernmost portion of panther range the disease could effectively spread only to the
south. Finally, vaccination may have helped end the epizootic. Vaccination of free-
ranging panthers began in August 2003, and as of June 2005, 34 panthers have received
at least one inoculation. Six of these have died due to non-FeLV causes, therefore, based
on a population size of between 80 and 100, approximately 28 to 35% of the population
has received at least one inoculation. However, because vaccination efforts were targeted
at OKS and adjacent lands, the percentage vaccinated in these areas is much greater.
Using computer models, Lubkin et al. (1996) estimated that 23% to 73% of a population
with a FeLV prevalence of 10% must be effectively vaccinated to eliminate infection.
Kennedy-Stoskopf (1999) speculated "[t]he lack of antigen-positive animals and
absence of clustered clinical cases with FeLV-related diseases are evidence that the virus
is not maintained in [non-domestic felid] populations." However, the finding of five
antigenemic panthers over almost 2 yr is evidence that the disease had, at least
temporarily, become established in the Florida panther population. Small populations are
at greater risk of extinction due to infectious diseases than larger populations
(Berger, 1990). With the exception of the occasional dispersing male, all free-ranging
Florida panthers are part of a single contiguous population in south Florida. As such, the
population is at risk of a catastrophic disease outbreak. Also of concern is the apparent
increased FeLV susceptibility of panthers compared to domestic cats. Transmission
appears to be occurring despite low infection pressure (few exposure events) and host
maturity, and the progression of infection appears to be more rapid compared to domestic
cats. Finally, FeLV prevalence among free-ranging domestic cats in Florida is <4% (Lee
et al., 2002), and in a review of FeLV in domestic cats, Levy (1999) speculated that
"[t]rue 'outbreaks' of FeLV infection are unlikely to occur." However, the prevalence of
FeLV in panthers sampled in OKS went from 0% prior to 2002 to 45.5% between
November 2002, and June 2005. If the disease spreads to the core population the impact
could be devastating.
More research is needed to further elucidate the epizootiology of FeLV in panthers.
Western blot antibody tests are needed to confirm positive ELISA antibody tests. Further,
quantitative PCR may be used to estimate provirus burden in panthers. This technique
may be more sensitive than conventional PCR for detecting latent infections. (Hofmann-
Lehmann et al., 2001). In domestic cats latent infections may eventually be cleared.
Testing to determine if living PCR-positive panthers converted to negative status should
be performed to determine the duration of the latent state. Finally, further research is
needed to determine the source of infection. Bobcats should be tested in the OKS region.
Domestic cats from this area should be tested as well, and virus recovered from infected
cats should be sequenced for comparison to strains isolated from panthers.
FLORIDA PANTHER/TEXAS PUMAS SAMPLED DURING THE STUDY PERIOD
Table A-1. Florida panthers and Texas pumas tested for feline leukemia virus (FeLV) antigen by ELISA 1 July 2002, to 5 June 2005.
Age Antibody Antibody ELISA Western
IDa Date Eventb SexC (yr) Locationd Areae ODf resultg antigenh Blot'
Table A-1. Continued.
Age Antibody Antibody ELISA Western
IDa Date Eventb Sex' (yr) Locationd Areae OD result' antigens Bloth
Table A-1. Continued.
Age Antibody Antibody ELISA Western
IDa Date Eventb Sex' (yr) Locationd Areae OD result' antigens Bloth
FP 120 4/8/2003 L F 3 BCNP-C South 0.193 N N N
FP 120 7/14/2004 L F 4 BCNP-C South 0.163 N N P
FP 121 12/2/2003 L F 2.5 SIR North 0.148 N N N
FP 122 1/30/2004 L F 2.25 OKS North 0.183 N P N
FP 122 2/13/2004 N F 2.25 OKS North P -
FP 123 2/2/2004 L M 3.5 OKS North 0.151 N P N
FP 123 3/15/2004 N M 3.6 OKS North P -
FP 124 2/13/2004 L F 3.5 BCNP-S South 0.155 N N P
FP 125 2/13/2004 L M 0.67 BCNP-S South 0.135 N N N
FP 126 2/13/2004 L M 0.67 BCNP-S South 0.155 N N N
FP 126k 5/28/2004 L M 0.96 BCNP-S South 0.163 N N
FP 127 2/16/2004 L M 2 BCNP-C South 0.149 N N P
FP 127k 3/29/2005 L M 3 BCNP-N South 0.196 N N P
FP 128 2/18/2004 L F 3.7 SIR North 0.171 N N N
FP 129 2/20/2004 L F 3 BCNP-C South 0.157 N N P
FP 130 3/4/2004 L M 0.8 OKS North 0.181 N N N
FP 130k 3/10/2005 L M 1.83 PL-Highlands Co. North 0.385 M N P
FP 131 3/10/2004 L M 5 FPNWR North 0.193 N N N
FP 132 3/17/2004 L M 3 OKS North 0.108 N N N
FP 132 7/22/2004 N M 3.33 OKS North 0.23 N P N
FP 133 11/18/2004 L M 4.5 BCNP-N North 0.214 N N P
FP 134 12/14/2004 L M 2.5 BCNP-N North 0.246 N N P
FP 135 12/17/2004 L M 1.7 FPNWR North 0.24 N N N
FP 136 1/13/2005 L F 5 BCNP-C South 0.207 N N N
FP 137 1/25/2005 L M 2.5 OKS North 0.239 N N N
FP 138 1/31/2005 L M 4 BCNP-C South 0.212 N N P
FP 139 3/31/2005 L M 2.83 OKS North 0.213 N N N
Table A-1. Continued.
Age Antibody Antibody ELISA Western
IDa Date Eventb Sex' (yr) Locationd Areae OD result' antigens Bloth
K 94 8/17/2004 N M 3 175 North-South N -
K 149 6/11/2003 L F 0.05 OKS North N -
K 150 6/11/2003 L M 0.05 OKS North N -
K 156 8/2/2004 N M 0.5 US41 South N -
K 175 2/10/2005 L M 0.04 BCNP-S South N -
K 176 2/10/2005 L M 0.04 BCNP-S South N -
K 178 3/7/2005 L M 0.05 OKS North N -
K 179 3/7/2005 L F 0.05 OKS North N -
K 180 3/21/2005 L F 0.05 FPNWR North N -
K 181 3/21/2005 L F 0.05 FPNWR North N -
TX 105 1/27/2003 L F 10.5 ENP South 0.161 N N P
TX 106 1/8/2003 L F 9.5 PSSF South 0.129 N N P
TX 108 11/18/2002 L F 10 ENP South 0.189 N N N
Table A-1. Continued.
Age Antibody Antibody ELISA Western
IDa Date Eventb Sex' (yr) Locationd Areae OD result' antigens Bloth
195 Flagler Co.
TABLE A-i LEGEND
aFP (Florida panther), TX (Texas puma), K (Florida panther previously handled as a
neonatal kitten), UCFP (Uncollared Florida panther).
bL (live-capture), N (necropsy)
cF (female), M (male).
dBCNP (Big Cypress National Preserve C [BCNP between 1-75 and US-41], N [BCNP
north of 1-75], S [BCNP south of US-41]), CR (County Road), CWMA (Crew Wildlife
Management Area), ENP (Everglades National Park), FPNWR (Florida Panther National
Wildlife Refuge), I (Interstate), OKS (Okaloacoochee Slough State Forest), PL (private
lands), PSSF (Picayune Strand State Forest), SIR (Big Cypress Seminole Indian
Reservation), SR (State Road), US (United States Road).
eN (panther range north of 1-75), S (panther range south of 1-75), N-S (killed by vehicular
collision on 1-75).
'OD (optical density).
gN (negative), L (low positive), M (medium positive), H (high positive).
hN (negative), P (positive).
'N (negative), P (positive), E (equivocal).
JTest results unsuitable due to severe autolysis.
kPreviously vaccinated for feline leukemia virus.
CASE REPORTS: ANTIGENEMIC FLORIDA PANTHERS
On 26 November 2002, a 4.5 yr-old female Florida panther was captured in OKS.
Capture was routine and the panther appeared healthy weighing 52.7 kg. Physical exam
was unremarkable and routine biomedical samples were collected. The panther was
vaccinated with Fel-O-Vax PCT, de-wormed, radio-instrumented and released.
Complete blood count revealed a mild non-regenerative anemia (28.4%), low
hemoglobin (9.2 g/dL), and lymphopenia (736/4l). Biochemical alterations were limited
to an elevated BUN (57 mg/dL), glucose (183 mg/dL), and creatinine phosphokinase
(609 U/L). FeLV ELISA antigen and FIV ELISA antibody (KELA and Western Blot)
were positive, although IFA of blood smears were inconclusive. Feline leukemia virus
was cultured from EDTA whole blood at the Ohio State University (OSU).
Radio-telemetry over the ensuing 5 mo indicated normal movements with a
minimum convex polygon (MCP) home range of 104.6 km2 (Shindle et al., 2003).
However, between 12 and 17 May 2003 movements became increasingly restricted. On
17 May 2003, the panther was located in a palmetto thicket and died at approximately
The panther was immediately collected, placed on ice, and transported to Disney's
Animal Kingdom the next day for complete necropsy. The panther had lost 20.4 kg since
capture; moderate SQ and abdominal fat were present although there was mild muscle
wasting. A copious red-tinged mucosy fluid drained from the nares. The right
submandibular lymph node exuded a bloody purulent yellow fluid. The tracheobronchial
lymph nodes were markedly enlarged and had a nodular appearance on cut section. A
frothy yellow mucoid material was observed at the tracheal bifurcation extending into the
distal airways. The lungs had a diffusely mottled red/black appearance with a nodular
texture which extended into the cut surface. The caudal lung lobes were most severely
affected. The thymus contained multifocal hemorrhages, and the pericardial sac contained
fibrin. The mesenteric lymph nodes were diffusely prominent. The stomach contained a
small amount of hog hair and mucus. The spleen was slightly enlarged and had a meaty
texture. The liver contained 1-3 mm multifocal tan foci. Each ovary contained two 3-5
mm diameter corpora lutea, and the uterus showed no evidence of previous pregnancy.
Histologically, the lungs contained nodular collections of alveoli containing dense
colonies of gram-negative bacteria intermixed with degenerative neutrophils, fibrin,
extravasated erythrocytes, and necrotic debris. There was necrosis of type I pneumocytes
and scattered hyperplasia of type II pneumocytes. Adjacent alveoli contained edema
fluid, macrophages, and neutrophils. Within the kidneys there was multifocal
mineralization of cortical tubules associated with necrosis of tubular epithelial cells. The
tubular epithelial cells contained a golden brown granular pigment. Throughout the
splenic parenchyma there was a mild increase of macrophages with mild hyperplasia of
white pulp. Megakaryocytes were scattered throughout the red pulp. Fibrin was seen in a
few splenic sinuses. Within the submandibular lymph node there was a focally extensive
area of necrosis infiltrated by large numbers of degenerate neutrophils. Other lymph
nodes showed evidence of multifocal cortical hyperplasia with sinuses containing
macrophages and lymphocytes. All other tissues appeared histologically normal.
Immunohistochemistry of spleen and lymph node was positive for p27 antigen.
Aerobic culture of the lung, liver, and submandibular lymph node resulted in pure
growth of E. coli. Viral isolation of the lung was negative.
Although IFA was inconclusive at capture, the persistence of antigenemia for >5
mo combined with clinical signs is consistent with persistent infection.
FP109 was initially captured and radio-instrumented as a 10 yr-old male 10
February 2002, in OKS. At capture he had injuries consistent with intraspecific
aggression. ELISA ag for FeLV at Cornell University was negative as was ELISA
antibody (HVL). Ten days later FP109 had to be recaptured to replace a defective radio-
collar and ELISA ag was again negative. The radio-collar failed several wk later.
FP109 was recaptured 24 January 2003, at 11 yr of age. At capture he appeared to
be in excellent condition but had a pronounced peripheral lymphadenopathy. Benign
hyperplasia was diagnosed from fine-needle aspirates of the popliteal lymph nodes.
Complete blood count revealed a moderate non-regenerative anemia (23.8%), low
hemoglobin (7.5 g/dL), and lymphopenia (490/4l). Rare C. felis organisms were seen on
blood smears. Serum biochemical abnormalities were minor with only an elevated
glucose (190 mg/dL) and decreased triglycerides (10 mg/dL). Feline leukemia virus
ELISA antigen and antibody tests were positive although IFA of blood smears were
inconclusive. Feline leukemia virus was cultured from EDTA whole blood at OSU.
Radio-telemetry indicated normal movements, but on 27 February 2003, FP109
was found dead. The panther had been dead for 2-3 days, and the carcass was severely
autolyzed, decomposed, and partially scavenged. Partial necropsy revealed puncture
wounds in the skin over the nasal bones with symmetrical crushing fractures of the nasal
bones. Injuries were consistent with ISA.
FP109 died before his true FeLV status could be determined. At capture in January
2003 he was likely in the early stages of infection. Given his high antibody OD, it is
possible he may have eventually cleared the infection. Negative ELISA antigen findings
in heart blood collected at necropsy supports this speculation; however, the sample was
extremely autolyzed and should be considered unreliable.
On 30 January 2004, a 2.25 yr-old female Florida panther was captured in OKS.
Capture was routine and the panther appeared in poor health weighing only 32.3 kg with
minimal SQ fat. Physical exam was otherwise normal except for a peripheral
lymphadenopathy. Routine biomedical samples were collected, and a SNAP test using
EDTA whole blood in the field was positive. The panther was vaccinated, de-wormed,
radio-instrumented, and released.
Complete blood count revealed a moderate non-regenerative anemia (22.5%), low
hemoglobin (7.2 g/dL), and monocytosis (1020/4l). Mild polychromasia, mild to
moderate anisocytosis, and 10 nucleated red blood cells/100 leukocytes were seen on
peripheral blood smear. Additionally, large immature mononuclear cells that occasionally
contained nucleoli were also seen; these findings were interpreted as an acute
lymphoblastic leukemia. Serum biochemical abnormalities included low cholesterol (62
mg/dL) and triglycerides (9 mg/dL). Abnormalities seen on urinalysis of free-catch urine
included 1+ blood, 3-10 WBC/HPF, 1-3 RBC/HPF, and 4+ bacteria/HPF. Specific
gravity was 1.009. ELISA antigen and IFA were positive. Virus was cultured from EDTA
blood at OSU.
Radio-telemetry over the ensuing 2 wk indicated normal movements and she
remained within the OKS area. Approximately 1 wk after capture, field sign indicated the
panther had killed and fed on a white-tailed deer (Odocoilius virginianus). However, on
13 February 2004, a mortality signal was detected, and her carcass was found in a
hammock in OKS. Time of death was approximately 0500 hrs.
The panther was immediately collected, placed on ice, and transported to Disney's
Animal Kingdom the next day for complete necropsy. At necropsy the panther was
approximately 15% dehydrated, in poor body condition, and had lost 8.9 kg since
capture. Mucus membranes and skeletal muscle were pale. Abdominal and SQ fat were
negligible, and there was evidence of serious fat atrophy. Adrenal glands were diffusely
enlarged. Peripheral lymph nodes were markedly enlarged.
Histologically the bone marrow was hypercellular with approximately 90% of the
marrow space occupied by hematopoietic cells. There was also a moderate increase in the
number of megakaryocytes. Erythroid precursor cells were decreased in number, and few
maturing erythroid cells were present. Myeloid cell lines were relatively increased in
number, and all stages of maturation were observed. Few lymphoid precursors and
mature lymphocytes were seen, and there was no marrow evidence of an acute leukemia.
There was no evidence of cortical follicle formation in examined lymph nodes.
Thymocytes were present in the thymus, but there was no evidence of cortical or
medullary architecture; intermixed among the thymocytes were macrophages containing
a bland golden brown pigment. Alveoli contained eosinophilic fluid and mildly increased
numbers of alveolar macrophages. Within the spleen, much of the red pulp was autolyzed
and there was scattered extramedullary hematopoiesis evident with few megakaryocytes
noted. Within the kidneys, scattered glomeruli were shrunken with markedly thickened
Bowman's capsules, collapse of the glomerular tufts, and replacement of the glomerular
tufts by fibrillar eosinophilic material. Rare cortical tubules were dilated. Scattered few
perivascular infiltrates of lymphocytes and plasma cells were present in the cortical
Rabies IFA was negative. Brain and heart were negative for CDV, pseudorabies
virus, flaviviruses, and alphaviruses by real-time and conventional PCR and viral culture.
Persistent infection was diagnosed based on positive ELISA antigen and IFA
results and clinical signs.
FP123, a 3.5 yr-old male, was captured 2 February 2004, in OKS. Capture was
routine and the panther appeared healthy weighing approximately 64 kg. Physical exam
was unremarkable and routine biomedical samples were collected. FeLV SNAP test using
EDTA whole blood was positive. The panther was vaccinated, dewormed, radio-
instrumented and released.
Complete blood count abnormalities were limited to a lymphopenia (884/kl),
although a significant percentage (11%) of the differential contained large, apparently
immature, mononuclear cells that occasionally contained nucleoli. These findings were
interpreted as an acute lymphoblastic leukemia. Serum biochemical abnormalities were
suggestive of dehydration and recent feeding (BUN 59 mg/dL, sodium 161 mEq/L,
BUN/creatinine ratio 39, triglycerides 222 mg/dL, and calculated osmolality 336
mOsm/L). FeLV ELISA antigen at Cornell Diagnostic Laboratory and IFA of blood
smears at the National Veterinary Laboratory were positive.
FP123 had a large home range (164.5 km2) traveling at least 10 km to the south
(Shindle et al., 2004). However, within 6 wk of capture, FP123 was found dead 17 March
2004, in OKS following detection of a mortality signal. The carcass was severely
autolyzed and decomposed; date of death was believed to have been 15-16 March. FP132
was captured the same day within 400 m of FP123. Acute injuries on FP132 were
consistent with ISA.
FP123 was completely necropsied at Disney's Animal Kingdom. No gross
abnormalities were noted although the carcass was severely autolysed.
On 17 March 2004, the carcass of FP123, a FeLV positive male, was recovered in
OKS. External injuries indicated the cause of death to be intraspecific aggression, and the
panther appeared to have been dead for approximately 24-48 hrs. Within 400 m of the
carcass, a freshly killed white-tailed deer was discovered. The dogs were released and
FP132, a 3 yr-old male, was captured. Capture was routine and the panther appeared
healthy weighing 66.3 kg. Two acute puncture wounds over the right shoulder,
presumably bite wounds, were seen on physical examination. Minor lacerations
consistent with claw marks were also seen. Circumstantial evidence was consistent with
FP132 as the cause of death for FP123. Routine biomedical samples were collected and a
SNAP test using EDTA whole blood in the field was negative. The panther was
vaccinated (including 2 ml Fel-O-Vax LvK), de-wormed, radio-instrumented, and
Complete blood count and serum chemistry were unremarkable. Repeat FeLV
ELISA antigen test was negative.
Radio-telemetry over the ensuing 4 mo indicated normal movements with a home
range of approximately 197.4 km2 (Shindle et al., 2004). FP132 was treed and boostered
with 2 ml Fel-O-Vax LvK on 12 April 2004. However, detectable movement based on
radio telemetry ceased between 14 and 21 July 2004. On 20 and 21 July, biologists
investigated and were able to approach to within 5 m of FP132 in thick brush before he
would move ahead. He appeared alert and healthy but lethargic. On 22 July, the panther
was located in a palmetto thicket and appeared to be in respiratory distress. He died at
approximately 1000 hr.
Within 30 min of death, whole blood was collected by dissection of the brachial
artery and aspiration with a needle and syringe. Blood was placed in serum separator and
EDTA tubes, and blood smears were made from cells (EDTA). The panther was
transported from the field and was on ice within 3 hr of death. Necropsy was performed
at Disney's Animal Kingdom. At necropsy the panther was approximately 10%
dehydrated and had lost over 13 kg since capture. There was moderate muscle wasting
although moderate to heavy SQ and abdominal fat was present. A 15x15 mm
pedunculated cutaneous mass was present over the left nasomaxillary region. Mucus
membranes were icteric and pale, and a copious red-tinged fluid drained from the nares.
Several healing puncture wounds and abrasions were noted in multiple sites.
Gross examination revealed a large abscess occupying the subcutis over the lateral
aspect of the right quadriceps muscle. The abscess measured 29x17 cm and had a
variable depth of 5-10 cm. The abscess contained several liters of tan cloudy fluid.
Skeletal muscle was pale. Lungs were diffusely dark red and firmer than expected. On cut
section numerous 1-5 mm tan foci were observed in all lung lobes although the left
cranial and medial lobes were most severely affected. Sections of lung tissue from these
lobes did not float in formalin. The liver was pale and friable. Peripheral lymph nodes
were not significantly enlarged, but mesenteric lymph nodes were larger than expected.
Aerobic cultures were taken of the abscess and lungs resulting in heavy growth of 3-
hemolytic Streptococcus sp.
Histologically, the skeletal muscle beneath the abscess was covered by a thick band
of mixed inflammatory cells representing the margin of the abscess. The superficial
aspect was composed of large numbers of degenerate neutrophils subtended by mixed
macrophages, lymphocytes, and plasma cells as well as immature fibroblasts and
connective tissue markedly expanded by edema. Numerous colonies of large bacterial
cocci were present on the superficial aspect of the lesion. Multifocally throughout the
lung, large dense colonies of bacteria and associated inflammatory cell aggregates
effaced the pulmonary architecture. Smaller bacterial colonies were also common in
airways. Large numbers of degenerate neutrophils and alveolar macrophages were
present in association with bacterial colonies and within the adjacent parenchyma. There
was necrosis of alveolar epithelium with multifocal type II pneumocyte hyperplasia.
Alveoli often contained strands of fibrin and edema fluid. Large areas of necrosis and
hemorrhage were also present. Numerous small (approx 50um diameter) objects
resembling trematode eggs were scattered throughout the liver. These eggs were bounded
by a refractile rim with the central core composed of granular basophilic material.
Numerous eggs were mineralized, degenerate, and were associated with small numbers of
macrophages. Within the thymus there was a loss of architecture and replacement by
abundant adipose tissue. There was also loss of cortico-medullary demarcation with
lymphocytes remaining in a loosely arranged fibrovascular stroma. Scattered cystic
structures were present and were presumed to be Hassals corpuscles. Also present within
the thymus were small mineralized structures which also represented calcified Hassals
corpuscles. Increased numbers of large macrophages containing pale brown cytoplasmic
pigment were present in the thymic parenchyma. Sections of bone marrow were
hypercellular with approximately 100% of the marrow space occupied by hematopoietic
cells. Megakaryocytes were present in normal to mildly increased numbers.
Serum biochemical abnormalities were consistent with hepatic failure (total
bilirubin 5.8 U/L, ALT 455 U/L, and AST 728 U/L) and pre-renal azotemia and/or renal
failure (BUN 63 mg/dL, creatinine 2.8 mg/dL). Other evidence of renal failure included a
severe hyperkalemia (9.7 mEq/L), hyperphosphotemia (19.1 mEq/L), hypermagnesemia,
and calculated osmolality (338 mOsm/L). These findings were likely a combination of
post-mortem artifact (potassium released from platelets), tissue necrosis, metabolic
acidosis, and dehydration. Hypoglycemia (61 mg/dL) was likely the result of septicemia.
FeLV SNAP test of serum and aqueous humor, and IFA of blood smears were
positive. Immunohistochemistry of spleen and lymph node were positive for p27 antigen.
ELISA antigen of serum at Antech Diagnostics was negative, but this is believed to be an
erroneous result. Virus was cultured at OSU.
Table B-1. Selected hematological and serum biochemical values for Florida panthers
testing positive for feline leukemia virus (FeLV) antigen by ELISA 1 July
2002 to 5 June 2005.
FIVe western blot
FeLV IFA blood smear
Blood urea nitrogen
White blood cells
Red blood cells (RBC)
Mean cell volume
Mean cell hemoglobin (MCH)
Nucleated RBC's (NRBC)
FP 109 FP 115 FP 122 FP 123 FP 132
M F F M M
1/24/2003 11/26/2002 1/30/2004 2/2/2004 722/2004
11 45 225 4 3
N P N N N
P P P P P
N N P P
190 183 115 120 61
22 57 24 59 63
20 21 24 15 28
80 66 65 74 55
34 32 33 36 27
01 02 03 03 58
7 12 8 7 4
45 36 35 62 455
24 36 28 68 728
104 93 103 98 102
36 48 50 52 191
46 34 32 38 28
223 609 244 470 223
75 92 72 132 -
23 8 284 225 425
49 92 85 52 -
491 685 418 875
48 41 54 49
153 134 172 151 -
315 324 320 31 1 -
147 236 185 350
4165 7360 4845 3848
85 80 57 74
0 0 0 0
0 0 0 0
490 736 2550 884
10 8 30 17
245 368 1020 416
5 4 12 8
0 736 85 52
0 8 1 1
0 0 0 0
0 0 0 0
0 10 0 -
13 16 13 02 02
P N N N
1 84 (0 54)
7 35 (0 67)
3 70 (0 36)
35 4 (386)
60 2 (35 0)
73 4 (77 8)
9 92 (0 66)
515 6 (415 1)
1221 (1 70)
X103/ 1 12 19(3 01)
X106/1l 7635 (1 033)
fl 47 29 (2 89)
Pg 1607(1 41)
g/dl 34 08 (3 26)
X103/1l 4026(131 5)
% WBC's 643(143)
% WBC's 28 8 (145)
% WBC's 3 2 (2 6)
% WBC's 3 4 (2 2)
% WBC's 0 89 (0 57)
'100WBC's 1 5 (1 0)
a ELISA (Enzyme-linked immunosorbent assay).
b Normal values for panthers (Dunbar et al., 1997).
c SD (standard deviation).
d OKS (Okaloacoochee Slough).
SFIV (feline immunodeficiency virus).
fIFA (immunofluorescent assay).
LIST OF REFERENCES
BARONE, M. A., M. E. ROELKE, J. HOWARD, J. L. BROWN, A. E. ANDERSON,
AND D. E. WILDT. 1994. Reproductive characteristics of male Florida panthers:
Comparative studies from Florida, Texas, Colorado, Latin America, and North
American zoos. Journal of Mammalogy 75: 150-162.
BEEBE, A. M., T. G. FAITH, E. E. SPARGER, M. TORTEN, N. C. PEDERSEN, AND
S. DANDEKAR. 1994. Evaluation of in vivo and in vitro interactions of feline
immunodeficiency virus and feline leukemia virus. AIDS 8: 873-878.
BERGER, J. 1990. Persistence of different-sized populations: an empirical assessment
of rapid extinctions in bighorn sheep. Conservation Biology 4: 91-98.
BIEK, R., R. L. ZARNKE, C. GILLIN, M. WILD, J. R. SQUIRES, AND M. POSS.
2002. Serologic survey for viral and bacterial infections in western populations of
Canada lynx (Lynx canadensis). Journal of Wildlife Diseases 38: 840-845.
BRIGGS, M. B., AND R. L. OTT. 1986. Feline leukemia virus infection in a captive
cheetah and the clinical and antibody response of six captive cheetahs to
vaccination with a subunit feline leukemia virus vaccine. Journal of the American
Veterinary Medical Association 189: 1197-1199.
CITINO, S. B. 1986. Transient FeLV viremia in a clouded leopard. Journal of Zoo and
Wildlife Medicine 17: 5-7.
COURCHAMP, F., C. SUPPO, E. FROMONT, AND C. BOULOUX. 1997. Dynamics
of two feline retroviruses (FIV and FeLV) within one population of cats.
Proceedings of the Royal Society of London B 264: 785-794.
CUNNINGHAM, M. W., M. R. DUNBAR, C. D. BUERGELT, B. L. HOMER, S. K.
TAYLOR, R. KING, S. B. CITINO, C. GLASS, AND M. E. ROELKE-PARKER.
1999. Atrial septal defects in Florida panthers. Journal of Wildlife Diseases
DANIELS, M. J., D. BALHARRY, D. HIRST, A. C. KITCHENER, AND R. J.
ASPINALL. 1998. Morphological and pelage characteristics of wild living cats in
Scotland: implications for defining the 'wildcat'. Journal of Zoology (London) 244:
,M. C. GOLDER, 0. JARRETT, AND D. W. MACDONALD. 1999. Feline
viruses in wildcats in Scotland. Journal of Wildlife Diseases 35: 121-124.
DUNBAR, M. R. 1994. Florida panther biomedical investigation, final performance
report. Endangered species project E-l-11 7506, Florida Game and Fresh Water
Fish Commission, Tallahassee, Florida, 51 pp.
S1995. Florida panther biomedical investigation, annual performance report.
Endangered species project E-l-11 7506, Florida Game and Fresh Water Fish
Commission, Tallahassee, Florida, 20 pp.
,P. NOL, AND S. B. LINDA. 1997. Hematologic and serum biochemical
reference intervals for Florida panthers. Journal of Wildlife Diseases 33:783-789.
CLAUDIA, F., J. L. CATAO-DIAS, G. BAY, E.L. DURIGON, R. S. P. JORGE, C. M.
LEUTENEGGER, H. LUTZ, AND R. HOFMANN-LEHMANN. (In press) Journal
of Wildlife Diseases.
FLYNN, J. N., S. P. DUNHAM, V. WATSON, AND O. JARRETT. 2002. Longitudinal
analysis of feline leukemia virus-specific cytotoxic T lymphocytes: correlation with
recovery from infection. Journal of Virology 76: 2306-2315.
FRANCIS, D. P., M. ESSEX, AND W. D. HARDY. 1977. Excretion of feline
leukaemia virus by naturally infected pets. Nature 269: 252-254.
AND D. GAYZAGIAN. 1979. Feline-leukemia virus: Survival under home
and laboratory conditions. Journal of Clinical Microbiology 9: 154-156.
FROMONT, E., M. ARTOIS, M. LANGLAIS, F. COURCHAMP, AND D. PONTIER.
1997. Modeling the feline leukemia virus (FeLV) in natural populations of cats
(Felis catus). Theoretical Population Biology 52: 60-70.
AND D. PONTIER. 1998a. Epidemiology of feline leukemia virus (FeLV)
and structure of domestic cat populations. Journal of Wildlife Management 62:
D. PONTIER, AND M. LANGLAIS. 1998b. Dynamics of a feline retrovirus
(FeLV) in host populations with variable spatial structure. Proceedings of the Royal
Society of London B 265: 1097-1104.
A. SAGER, F. LEGER, F. BOURGUEMEISTER, E. JOUQUELET, P. STAHL,
D. PONTIER, AND M. ARTOIS. 2000. Prevalence and pathogenicity of
retroviruses in wildcats in France. Veterinary Record 146: 317-319.
D. PONTIER, AND M. LANGLAIS. 2003. Disease propagation in connected
host populations with density-dependent dynamics: the case of the Feline leukemia
virus. Journal of Theoretical Biology 223: 465-475.
GERSTMAN, B. 1985. The epizootiology of feline leukemia virus infection and its
associated diseases. Compendium for Continuing Education 7: 766-774.
GRANT, C. K., M. ESSEX, M. B. GARDNER, W. D. HARDY. 1980. Natural feline
leukemia virus infection and the immune response of cats of different ages. Cancer
Research 40: 823-829.
GRINDEM, C. B., W. T. CORBETT, B. E. AMMERMAN, AND M. T. TOMKINS.
1989. Seroepidemiologic survey of feline immunodeficiency virus infection in cats
of Wake County, North Carolina. Journal of the American Veterinary Medical
Association 194: 226-228.
HARDY, W. D., JR. 1973. Horizontal transmission of feline leukaemia virus. Nature
1980a. Feline leukemia virus diseases. In Feline leukemia virus, W. D. HARDY,
JR., M. ESSEX, AND A. MCCLELLAND (eds.). Elsevier/North-Holland, New
York, New York, pp 3-31.
1980b. The virology, immunology and epidemiology of the feline leukemia virus.
In Feline leukemia virus, W. D. HARDY, JR., M. ESSEX, AND A.
MCCLELLAND (eds.). Elsevier/North-Holland, New York, New York, pp 33-78.
,L. J. OLD, P. W. HESS, M. ESSEX, AND S. M. COTTER. 1973. Horizontal
transmission of feline leukemia virus. Nature 244: 266-269.
,P. W. HESS, E. G. MACEWEN, A. J. MCCLELLAND, E. E. ZUCKERMAN, M.
ESSEX, S. M. COTTER, AND O. JARRETT. 1976. Biology of feline leukemia
virus in the natural environment. Cancer Research 36: 582-588.
HARTMANN, K. 2005. Pathogenesis of FeLV. In Clinical advances: A supplement to
compendium on continuing education for the practicing veterinarian 27(2A): pp. 8-
HIETALA, S. K., AND I. A. GARDNER. 1999. Validity of using diagnostic tests that
are approved for use in domestic animals for nondomestic species. In Zoo and wild
animal medicine, M. E. FOWLER AND R. E. MILLER (eds.). W. B. Saunders
Co., Philadelphia, Pennsylvania, pp. 55-58.
HOFMANN-LEHMANN, R., E. HOLZNAGEL, A. AUBERT, P. OSSENT, M.
REINACHER, AND H. LUTZ. 1995. Recombinant FeLV vaccine: long-term
protection and effect on course and outcome of FIV infection. Veterinary
Immunology and Immunopathology 46: 127-137.
,D. FEHR, M. GROB, M. ELGIZOLI, C. PACKER, J. S. MARTENSON, S. J.
O'BRIEN, H. LUTZ. 1996. Prevalence of antibodies to feline parvovirus,
calicivirus, herpesvirus, coronavirus, and immunodeficiency virus and of feline
leukemia virus antigen and the interrelationship of these viral infections in free-
ranging lions in East Africa. Clinical and Diagnostic Laboratory Immunology 3:
E. HOLZNAGEL, P. OSSENT, AND H. LUTZ. 1997. Parameters of disease
progression in long-term experimental feline retrovirus (feline immunodeficiency
virus and feline leukemia virus) infections: hematology, clinical chemistry, and
lymphocyte subsets. Clinical and Diagnostic Laboratory Immunology 4: 33-42.
J. B. HUDER, S. GRUBER, F. BORETTI, B. SIGRIST, AND H. LUTZ. 2001.
Feline leukemia provirus load during the course of experimental infection and in
naturally infected cats. Journal of General Virology 82:1589-1596.
HOOVER, E. A., AND J. I. MULLINS. 1991. Feline leukemia virus infection and
disease. Journal of American Veterinary Medical Association 199: 1287-1297.
R. G. OLSEN, W. D. HARDY, JR., J. P. SCHALLER, AND L. E. MATCHES.
1976. Feline leukemia virus infection: age-related variation in response of cats to
experimental infection. Journal of the National Cancer Institute 57: 365-369.
J. L. ROJKO, AND R. G. OLSEN. 1980. Factors influencing host resistance to
feline leukemia virus. In Feline leukemia, R. G. OLSEN (ed.). CRC Press, Boca
Raton, Florida, pp. 69-76.
JACOBSON, R. H., AND N. A. LOPEZ. 1991. Comparative study of diagnostic testing
for feline leukemia virus infection. Journal of Veterinary Medical Association 199:
JARRETT, O. 1983. Recent advances in the epidemiology of feline leukaemia virus.
Veterinary Annual 23: 287-293.
H. M. LAIRD, AND D. HAY. 1973. Determinants of host range of feline
leukaemia viruses. Journal of General Virology 20: 169-175.
W. D. HARDY, JR., M. C. GOLDER, AND D. HAY. 1978. The frequency of
occurrence of feline leukemia virus subgroups in cats. International Journal of
Cancer 21: 334-337.
M. C. GOLDER, AND K. WEIJER. 1982. A comparison of three methods of
feline leukaemia virus diagnosis. The Veterinary Record 110: 325-328.
JESSUP, D. A., C. PETTAN, L. J. LOWENSTINE, AND N. C. PEDERSON. 1993.
Feline leukemia virus infection and renal spirochetosis in a free-ranging cougar
(Felis concolor). Journal of Zoo and Wildlife Medicine 24: 73-79.
KENNEDY-STOSKOPF, S. 1999. Emerging viral infections in large cats. In Zoo and
wild animal medicine, M. E. FOWLER AND R. E. MILLER (eds.). W. B.
Saunders Co., Philadelphia, Pennsylvania, pp. 401-410.
LAND, E. D., D. R. GARMAN, AND G. A. HOLT. 1998. Monitoring female Florida
panthers via cellular telephone. Wildlife Society Bulletin 26: 29-31.
,M. CUNNINGHAM, R. MCBRIDE, D. SHINDLE, AND M. LOTZ. 2002.
Florida panther genetic restoration and management: annual report. Florida Fish
and Wildlife Conservation Commission, Tallahassee, Florida, 111 pp.
LEE, I. T., J. K. LEVY, S. P GORMAN, P. C. CRAWFORD, AND M. R. SLATER.
2002. Prevalence of feline leukemia virus infection and serum antibodies against
feline immunodeficiency virus in unowned free-roaming cats. Journal of the
American Veterinary Medical Association 220:620-622.
LEVY, J. K. 1999. FeLV and non-neoplastic FeLV-related disease. In Textbook of
veterinary internal medicine, S. J. ETTINGER AND E. C. FELDMAN (eds.). W.
B. Saunders Co., Philadelphia, Pennsylvania, pp. 424-432.
S2005. Epidemiology, transmissibility, and risk assessment in FeLV. In Clinical
advances: A supplement to compendium on continuing education for the practicing
veterinarian 27(2A): pp. 4-7.
AND C. CRAWFORD. 2005. Feline leukemia virus. In Textbook of veterinary
internal medicine, S. J. ETTINGER AND E. C. FELDMAN (eds.). W. B. Saunders
Co., Philadelphia, Pennsylvania, pp. 653-659.
LOPEZ, N. 1988. Panther study provides new insight into FeLV tests. In Feline health
topics pp: 5-8.
,AND R. H. JACOBSON. 1989. False-positive reactions associated with anti-
mouse activity in serotests for feline leukemia virus antigen. Journal of Veterinary
Medical Association 195: 741-746.
LUBKIN, S. R., J. ROMATOWSKI, M. ZHU, P. M. KULESA, AND K. A. J. WHITE.
1996. Evaluation of feline leukemia virus control measures. Journal of Theoretical
LUTZ, H., N. PEDERSEN, J. HIGGINS, C. W. HARRIS, AND G. H. THEILEN.
1980a. Quantitation of p27 in the serum of cats during natural infection with feline
leukemia virus. In Feline leukemia virus, W. D. HARDY, M. ESSEX, AND A. J.
MCCLELLAND (eds.). Elsevier/North-Holland, New York, New York, pp. 497-
U. HUBSCHER, F. A. TROY, AND G. H. THEILEN. 1980b.
Humoral immune reactivity to feline leukemia virus and associated antigens in cats
naturally infected with feline leukemia virus. Cancer Research 40: 3642-3651.
MAEHR, D. S. 1997. The Florida panther: Life and death of a vanishing carnivore.
Island Press, Washington, D.C., 261 pp.
MANDEL, M. P., J. R. Stephenson, W. D. Hardy, Jr., and M. Essex. 1979. Endogenous
RD-114 virus of cats: absence of antibodies to RD-114 envelope antigens in cats
naturally exposed to the feline leukemia virus. Infection and Immunology 24: 282-
MANSFIELD, K. G., AND E. D. LAND. 2002. Cryptorchidism in Florida panthers:
prevalence, features, and influence of genetic variation. Journal of Wildlife
Diseases 38: 693-698.
MARKER, L., L. MUNSON, P. A. BASSON, AND S. QUACKENBUSH. 2003.
Multicentric T-cell lymphoma associated with feline leukemia virus infection in a
captive Namibian cheetah (Acinonyx ubatus). Journal of Wildlife Diseases 39:
MCBRIDE, R. 2003. The documented panther population (DPP) and its current
distribution from July 1, 2002 to June 30, 2003. Livestock Protection Company,
Alpine, Texas, 11 pp.
MCCLELLAND, A. J., W. D. HARDY, AND E. E. ZUCKERMAN. 1980. Prognosis of
healthy feline leukemia virus infected cats. In Feline leukemia virus, W. D.
HARDY, M. ESSEX, AND A. J. MCCLELLAND (eds.). Elsevier/North-Holland,
New York, New York, pp. 121-126.
MCCOWN, J. W., D. S. MAEHR, AND J. ROBOSKI. 1990. A portable cushion as a
wildlife capture aid. Wildlife Society Bulletin 18: 34-36.
MCMICHAEL, J. C., S. STIERS, AND S. COFFIN. 1986. Prevalence of feline
leukemia virus infection among adult cats at an animal control center: association
of viremia with phenotype and season. American Journal of Veterinary Research
MERIC, S. M. 1984. Suspected feline leukemia virus infection and pancytopenia in a
western cougar. Journal of the American Veterinary Medical Association 185:
MIYAZAWA, T. 2002. Infections of feline and feline immunodeficiency virus.
Frontiers in Bioscience 7: 504-518.
,Y. IKEDA, K. MAEDA, T. HORIMOTO, Y. TOHYA, M. MOCHIZUKI, D. VU.
G. D. VU, D. X. CU, K. ONO, E. TAKAHASHI, AND T. MIKAMI. 1997.
Seroepidemiological survey of feline retrovirus infections in domestic and leopard
cats in northern Vietnam in 1997. Journal of Veterinary Medical Science 60: 1273-
MOCHIZUKI, M., M. AKUZAWA, AND H. NAGATOMO. 1990. Serological survey
of the Iriomote cat (Felis iriomotensis) in Japan. Journal of Wildlife Diseases 26:
MUNSON, L., L. MARKER, E. DUBOVI, J. A. SPENCER, J. F. EVERMANN, AND S.
J. O'BRIEN. 2004. Serosurvey of viral infections in free-ranging Namibian
cheetahs (Acinonyx jubatas). Journal of Wildlife Diseases 40: 23-31.
NAKATA, R., T. MIYAZAWA, Y-S. SHIN, R. WATANABE, T. MIKAMI, AND Y.
MATSUURA. 2003. Reevaluation of host ranges of feline leukemia virus
subgroups. Microbes and Infection 5: 947-950.
NEWMAN, A., M. BUSH, D. E. WILDT, D. VAN DAM, M. T. FRANKENHUIS, L.
SIMMONS, L. PHILLIPS, AND S. J. O'BRIEN. 1985. Biochemical genetic
variation in eight endangered or threatened felid species. Journal of Mammalogy
NOWAK, R. M., AND R. MCBRIDE. 1973. Status survey of the Florida panther.
Reprinted from the World Wildlife Fund Yearbook, 1973-1974. In Proceedings of
the Florida panther conference. Florida Audubon Society and Florida Game and
Fresh Water Fish Commission, Orlando, Florida, p. 118.
O'BRIEN, S. J., M. E. ROELKE, N. YUHKI, K. W. RICHARDS, W. E. JOHNSON, W.
L. FRANKLIN, A. E. ANDERSON, O. L. BASS, JR., R. C. BELDEN, AND J. S.
MARTENSON. 1990. Genetic introgression within the Florida panther Felis
concolor coryi. National Geographic Research 6: 485-494.
OLMSTEAD, R. A., R. LANGLEY, M. E. ROELKE, R. M. GOEKEN, D. ADGER-
JOHNSON, J. P. GOFF, J. P. ALBERT, C. PACKER, M. K. LAURENSON, T. M.
CARO, L. SCHEEPERS, D. E. WILDT, M. BUSH, J. S. MARTENSON, AND S.
J. O'BRIEN. 1992. Worldwide prevalence of lentivirus infection in wild feline
species: epidemiologic and phylogenetic aspects. Journal of Virology 66: 6008-
OSOFSKY, S. A., K. J. HIRSCH, E. E. ZUCKERMAN, AND W. D. HARDY, JR..
1996. Feline lentivirus and feline oncovirus status of free-ranging lions (Panthera
leo), leopards (Panthera pardus), and cheetahs (Acinonyx jubatus) in Botswana: a
regional perspective. Journal of Zoo and Wildlife Medicine 27: 453-467.
OSTROWSKI, S., M. VAN VUUREN, D. M. LENAIN, AND A. DURAND. 2003. A
serologic survey of wild felids from central west Saudi Arabia. Journal of Wildlife
PAUL-MURPHY, J., T. WORK, D. HUNTER, E. MCFIE, AND D. FJELLINE. 1994.
Serologic survey and serum biochemical reference ranges of the free-ranging
mountain lion (Felis concolor) in California. Journal of Wildlife Diseases 30: 205-
PEDERSEN N. C., S. M. MERIC, L. J. JOHNSON, S. P. PLUCKER, AND G. H.
THEILEN. 1984. The clinical significance of latent feline leukemia virus infection
and cats. Feline Practice 14: 32-48.
,M. TORTEN, B. RIDEOUT, E. SPARGER, T. TONACHINI, P. A. LUCIW, C.
ACKLEY, N. LEVY, AND J. YAMAMOTO. 1990. Feline leukemia virus
infection as a potentiating cofactor for the primary and secondary stages of
experimentally induced feline immunodeficiency virus infection. Journal of
Virology 64: 598-606.
RAMOS-VARA, J. A., M. KIUPEL, AND M. A. MILLER. 2002. Diagnostic
immunohistochemistry of infectious diseases in dogs and cats. Journal of
Histotechnology 25: 201-214.
RASHEED, S., AND M. B. GARDNER. 1981. Isolation of feline leukemia virus from a
leopard cat cell line and search for retrovirus in wild felidae. Journal of the
National Cancer Institute 67: 929-933.
REINACHER, M. 1989. Diseases associated with spontaneous feline leukemia virus
(FeLV) infection in cats. Veterinary Immunology and Immunopathology 21: 85-95.
,G. WITTMER, H. KOBERSTEIN, AND K. FAILING. 1995. Untersuchungen
zur bedeutung der FeLV- infection fur erkrankungen bei sektionskatzen. Berliner
und Munchener tierarztliche Wochenschrift. 108: 58-60.
RICKARD, L. G., AND W. J. FOREYT. 1992. Gastrointestinal parasites of cougars
(Felis concolor) in Washington and the first report of Ollulanus tricuspis in a
sylvatic felid from North America. Journal of Wildlife Diseases 28: 130133.
RIGBY, M. A., J. L. ROJKO, M. A. STEWART, G. J. KOCHIBA, C. M. CHENEY, L.
J. REZANKA, L. E. MATCHES, J. R. HARTKE, O. JARRETT, AND J. C. NEIL.
1992. Partial dissociation of subgroup C phenotype and in vivo behaviour in feline
leukaemia viruses with chimeric envelope genes. Journal of General Virology 73:
RILEY, S. P. D., J. FOLEY, AND B. CHOMEL. 2004. Exposure to feline and canine
pathogens in bobcats and gray foxes in urban and rural zones of a National Park in
California. Journal of Wildlife Diseases 40: 11-22.
ROELKE, M. E. 1990. Florida panther biomedical investigation, final performance
report. Endangered species project E-l-11 7506. Florida Game and Fresh Water
Fish Commission, Tallahassee, Florida, 178 pp.
,J. S. MARTENSON, AND S. J. O'BRIEN. 1993a. The consequences of
demographic reduction and genetic depletion in the endangered Florida panther.
Current Biology 3: 340-350.
,D. J. FORRESTER, E. R. JACOBSON, G. V. KOLIAS, F. W. SCOTT, M. C.
BARR, J. F. EVERMANN, AND E. C. PIRTLE. 1993b. Seroprevalence of
infectious disease agents in free-ranging Florida panthers (Felis concolor coryi).
Journal of Wildlife Diseases 29: 36-49.
ROGERSON, P., W. JARRETT, AND L. MACKEY. 1975. Epidemiological studies on
feline leukemia virus infection. International Journal of Cancer 15: 781-785.
ROJKO, J. L., AND G. J. KUCIBA. 1991. Pathogenesis of infection by the feline
leukemia virus. Journal of the American Veterinary Medical Association 199:
,E. A. HOOVER, L. E. MATCHES, R. G. OLSEN, AND J. P. SCHALLER. 1979.
Pathogenesis of experimental feline leukemia virus infection. Journal of the
National Cancer Institute 63: 759-768.
S. L. QUACKENBUSH, AND R. G. OLSEN. 1982. Reactivation of latent
feline leukaemia virus infection. Nature 298: 385-389.
ROTSTEIN, D. S., R. THOMAS, K. HELMICK, S. B. CITINO, S. K., TAYLOR, AND
M. R. DUNBAR. 1999. Dermatophyte infections in free-ranging Florida panthers
(Felis concolor coryi). Journal of Zoo and Wildlife Medicine 30: 281-284.
RYSER-DEGIORIS, M. P., R. HOFFMANN-LEHMANN, C. M. LEUTENEGGER, C.
HARD AF SEGERSTAD, T. MONER, R. MATTSSON, AND H. LUTZ. 2005.
Epizootiologic investigations of selected infectious disease agents in free-ranging
Eurasian lynx from Sweden. Journal of Wildlife Diseases 41: 58-66.
SARMA, P. S., AND T. LOG. 1973. Subgroup classification of feline leukemia virus
and sarcoma virses by viral interference and neutralization tests. Virology 54: 160-
SCHMITT, A. C., D. REISCHAK, C. L. CAVLAC, C. H. L. MONTFORTE, F. T.
COUTO, A. B. P. F. ALMEIDA, D. G. G. SANTOS, L. SOUZA, C. ALVES, K.
VECCHI. 2003. Infegcco pelos virus da leukemia feline e da peritonite infecciosa
feline em felideo selvagem de vida livre e cativeiro da regido do Pantanal
matogrossense. Acta Scientiae Veterinariae 31: 185-188.
SEAL, U. S. 1994. A plan for genetic restoration and management of the Florida
panther (Felis concolor coryi). Report to the Florida Game and Fresh Water Fish
Commission, Conservation Breeding Specialist Group, SSC/IUCN. White Oak
Conservation Center, Yulee, Florida, 23 pp.
,AND R. LACY. 1989. Florida panther (Felis concolor coryi) viability analysis
and species survival plan. Report to the U.S. Fish and Wildlife Service. Captive
Breeding Specialist Group, SSC/IUCN. Apple Valley, Minnesota, 208 pp.
SHINDLE, D., D. LAND, K. CHARLTON, AND R. MCBRIDE. 2000. Florida panther
genetic restoration and management: annual report. Florida Fish and Wildlife
Conservation Commission, Tallahassee, Florida, 94 pp.
M. CUNNINGHAM, D. LAND, R. MCBRIDE, M. LOTZ, AND B. FERREE.
2003. Florida panther genetic restoration and management: Annual report. Florida
Fish and Wildlife Conservation Commission, Tallahassee, Florida, 111 pp.
D. LAND, M. W. CUNNINGHAM, M. LOTZ, AND B. FERREE. 2004. Florida
panther genetic restoration and management: Annual report. Florida Fish and
Wildlife Conservation Commission, Tallahassee, Florida, 102 pp.
SLEEMAN, J. M., J. M. KEANE, J. S. JOHNSON, R. J. BROWN, AND S. V. WOUDE.
2001. Feline leukemia virus in a captive bobcat. Journal of Wildlife Diseases 37:
TAYLOR, S. K., C. D. BUERGELT, M. E. ROELKE-PARKER, B. L. HOMER, AND
D. S. ROTSTEIN. 2002. Causes of mortality of free-ranging Florida panthers.
Journal of Wildlife Diseases 38: 107-114.
TORRES, A. N., C. K. MATHIASON, AND E. A. HOOVER. 2005. Re-examination of
feline leukemia virus: host relationships using real-time PCR. Virology 332: 272-
VANAS, J. 1976. The Florida panther in the Big Cypress Swamp and the role of the
Everglades Wonder Gardens in past and future captive breeding programs. In
Proceedings of the Florida panther conference. P. C. H. PRICHARD (ed.). Florida
Audubon Society and Florida Game and Fresh Water Fish Commission, Orlando,
Florida, pp. 109-111.
Mark Cunningham was born in Chicago, Illinois, June 16, 1966, and was raised in
Miami, Florida. He received his BA in biology from Florida State University in 1991 and
graduated from the University of Florida, College of Veterinary Medicine in 1998. He is
currently employed as the Division of Wildlife Research veterinarian for the Florida Fish
and Wildlife Conservation Commission.