365The Laboratory Rabbit, Guinea Pig, Hamster, and Other Rodents [603156]

365The Laboratory Rabbit, Guinea Pig, Hamster, and Other Rodents
DOI: © 2012 Elsevier Inc.
2012 10.1016/B978-0-12-380920-9.00014-614
INTRODUCTION
The major focus of this chapter is the naturally occur –
ring viral diseases of rabbits and hares although asymp –
tomatic infections are also discussed ( Tables 14.1 and
14.2). Some viral infections of rabbits have provided
fundamental information on basic mechanisms of
agent–host interrelationships, e.g., Myxoma virus , and
others have been useful as models for human diseases, e.g., Rotavirus . Although the principal emphasis is
on virus infections of domestic rabbits of the genus
Oryctolagus, naturally occurring and experimental infec –
tions of other rabbits and hares are also discussed.
In indoor facilities, it is unlikely that rabbits will
encounter any of these viruses. Incoming rabbits should
be obtained from sources free of these infections, how –
ever appropriate quarantine and screening measures
should be standard operating procedure. With outdoor Introduction 365
DNA Virus Infections 368
Poxvirus Infections 368
Myxoma virus 368
Rabbit (shope) fibroma virus 373
Hare Fibroma Virus 376
Rabbitpox 376
Herpesvirus Infections 379
Leporid Herpesvirus 1 379
Leporid Herpesvirus 2 380
Leporid Herpesvirus 4 381
Human herpesvirus 1 (HHV-1) 382
Papilloma and Polyoma Virus Infections 382
Cottontail rabbit papillomavirus 382
Rabbit oral papillomavirus 384
Rabbit kidney vacuolating virus 385
Adenovirus Infections 385
Parvovirus Infections 386
RNA Virus Infections 386
Rotavirus Infections 386Rotavirus 386
Coronavirus Infections 389
Pleural Effusion Disease/Infectious
Cardiomyopathy Virus 389
Rabbit Enteric Coronavirus 391
Calicivirus Infections 392
Rabbit hemorrhagic disease virus (RHDV) 392
European brown hare syndrome virus
(EBHSV) 396
Rabbit Calicivirus (RCV) 397
Michigan Rabbit Calicivirus (MRCV) 398
Rabbit Vesivirus 399
Paramyxovirus Infections 399
Bunyavirus Infections 399
Togavirus Infections 400
Flavivirus Infections 400
Picobirnavirus Infections 401
Rabies Virus 401
References 401C H A P T E R
Viral Diseases
Thea Brabb and Ronald F . Di Giacomo
University of Washington, Seattle, WA, USA
O U T L I N E

II. RABBITS
14. VIRAL DISEASES
366
TABLE 14.1 DNA Virus Infections of Rabbits
Family/subfamily Genus Species/strain Host Geographic Distribution
Poxviridae Leporipoxvirus Myxoma virus Sylvilagus brasilensis South America
Sylvilagus bachmani North America
Oryctolagus cuniculus (wild and
domesticated)South America, North
America, Europe, Australia,
New Zealand
Rabbit (Shope) fibroma
virusSylvilagus floridanus North America
Hare fibroma virus Lepus europaeus Europe
Orthopoxvirus Rabbitpox Oryctolagus cuniculus (domesticated)aUnited States (laboratory
colonies), Holland
Herpesviridae/
GammaherpesvirinaeRhadinovirus Leporid herpesvirus 1
Leporid herpesvirus 2Sylvilagus floridanus
Oryctolagus cuniculus (domesticated)United States
England, United States
(laboratory colonies)
Herpesviridae/
AlphaherpesvirinaeSimplexvirus Leporid herpesvirus 4
Human herpesvirus 1Oryctolagus cuniculus (domesticated)
Oryctolagus cuniculus (domesticated)aNorth America
United States
Papillomaviridae Kappa-papillomavirus Cottontail rabbit
papillomavirusSylvilagus floridanus
Oryctolagus cuniculus (domesticated)United States
United States
Rabbit oral
papillomavirusOryctolagus cuniculus (domesticated) United States, Mexico, New
Zealand
Polyomaviridae Polyomavirus Rabbit kidney
vacuolating virusSylvilagus floridanus
Oryctolagus cuniculus (domesticated)United States
United States
Adenoviridae Mastadenovirus Adenovirus Oryctolagus cuniculus (domesticated) Hungary, Quebecb
Parvoviridae Parvovirus Lapine parvovirus Oryctolagus cuniculus (domesticated) Japan, United States,
Switzerlandb
aAberrant host.
bSerologic evidence only.
TABLE 14.2 RNA Virus Infections of Rabbits
Family Genus Species/strain Host Geographic Distribution
Reoviridae Rotavirus Rotavirus Oryctolagus cuniculus (domesticated) North America, Japan, Europe,
Sylvilagus floridanus Canadaa
Lepus americanusaCanadaa
Coronaviridae Coronavirus Pleura effusion disease Oryctolagus cuniculus (experimental) Europe, United States, Japan
Rabbit enteric coronavirus Oryctolagus cuniculus (domesticated) North America, Europe
Caliciviridae Lagovirus Rabbit hemorrhagic disease
virusOryctolagus cuniculus (wild and
domesticated)Europe, Asia, North Africa,
Middle East, Australia, New
Zealand, Mexico, United States
Lepus sp. China
European brown hare syndrome
virusLepus europaeus Europe, Argentina
(Continued )

367 INTRODUCTION
II. RABBITSLepus timidus Europe
Rabbit Calicivirus Oryctolagus cuniculus (wild and
domesticated)Europe, Australia
Michigan Rabbit Calicivirus Oryctolagus cuniculus (domesticated) United States
Vesivirus Not assigned Oryctolagus cuniculus (domesticated) United States
Paramyxoviridae Not assigned Rabbit syncytium virus Sylvilagus floridanus United States
Respivovirus Sendai-like virus Oryctolagus cuniculus (domesticated)aJapan
Bunyaviridae Orthobunyavirus California encephalitis virus Sylvilagus floridanusaNorth America
Lepus californicusaNorth America
Snowshoe hare Lepus americanus North America,
Tahyna Lepus europeausaEurope
Oryctolagus cuniculus (wild)aEurope
lnkoo Lepus timidusaFinland
Bunyamwera Cache valley Sylvilagus auduboniiaNorth America
Sylvilagus floridanus North America
Lepus californicusaNorth America
Tenshaw Sylvilagus acquaticusaUnited States
Sylvilagus palustrisaUnited States
Sylvilagus transitionalis United States
Northway Lepus americanus Alaska, California
Not assigned Silverwater Lepus americanus Canada
Togaviridae Alphavirus Western equine encephalitis
virusSylvilagus sp.a
Lepus californicusaNorth America
North America
Lepus americanusaNorth America
Oryctolagus cuniculus (domesticated) North America
Eastern equine encephalitis
virusLepus americanusaNorth America
Venezuelan (VEE) equine
encephalitis virusLepus americanusaNorth America
Sylvilagus floridanusaNorth America
Flaviviridae Flavivirus St. Louis encephalitis virus Lepus americanusaNorth America
West Nile virus Sylvilagus floridanus (experimental) North America
Powassan virus Lepus americanus North America, Russia
Picobirnaviridae Picobirnavirus Not assigned Oryctolagus cuniculus (domesticated) United States
Rhabdoviridae Lysavirus Rabies virus Sylvilatus floridanus United States
Oryctolagus cuniculus (domesticated) United States
aSerologic evidence only.TABLE 14.2 RNA Virus Infections of Rabbits (Continued)
Family Genus Species/strain Host Geographic Distribution

II. RABBITS
14. VIRAL DISEASES
368
facilities, appropriate measures should be undertaken,
as described in the “Control” section for each of the
viral diseases, to prevent spread of viruses from wild
rabbits in the surrounding environment. None of the
viral infections of rabbits are known to be of public
health importance as there are no reports of the defini –
tive spread of viruses from rabbits to humans.
The viral diseases of rabbits are discussed in an order
based on the taxonomic groups to which the viruses
belong and are independent of the order of importance
of the various diseases. The material is presented under
uniform subject headings, including history, etiology,
epidemiology, clinical signs, pathology, diagnosis, and
control. Control is interpreted broadly and includes both
prevention and eradication. This review uses the most
widely accepted viral terminology. The recommenda –
tions of the VIII International Committee on Taxonomy
of Viruses ( Fauquet et al., 2005 ) are followed, but well-
established common names for viruses are used when
appropriate. Rabbits and hares are referenced by com –
mon names as indicated in Mammals of the World, A
Checklist (Duff and Lawson, 2004 ).
DNA VIRUS INFECTIONS
Poxvirus Infections
Poxviruses cause several important diseases in
domestic and wild mammals and birds. Infection with
poxviruses usually results in relatively mild disease
involving the skin of infected animals, but generalized
and often fatal disease may also occur, as, for example,
in myxomatosis in rabbits. Close antigenic relationships
exist among many poxviruses derived from different
animal species. In spite of close antigenic relationships,
the poxviruses of rabbits which produce distinct disease
syndromes are discussed as separate entities.
Myxoma Virus
HISTORY
The disease myxomatosis, caused by Myxoma virus ,
was first recognized by Sanarelli (1898) in Uruguay
in 1896. European rabbits of the genus Oryctolagus,
acquired for antiserum production, developed a highly
fatal disease characterized by numerous mucinous skin
tumors. Sanarelli (1898) named the disease “infectious
myxomatosis of rabbits” and, since no microbial agents
were detected, proposed that the disease was caused by
a newly recognized group of infectious agents known
as “filterable viruses.” The virus which caused the first
known outbreak of myxomatosis is believed to have
originated from the Forest rabbit ( Sylvilagus brasiliensis )
in which the virus causes relatively mild disease.
Transmission from wild to domestic rabbits probably occurred by mosquitoes of the genus Aedes (Aragao,
1943; Fenner and Ratcliffe, 1965 ).
Myxomatosis spread to other countries of South
America where it occasionally causes sporadic out –
breaks in domesticated rabbits. In Chile, the disease is
considered endemic in the wild European rabbit popu –
lation ( Fenner and Ratcliffe, 1965 ). The disease was
first recognized in North America in 1928 when natural
outbreaks of a fatal disease of rabbits occurred in sev –
eral rabbit colonies near San Diego, California ( Kessel
et al., 1931 ). The virus which caused the first outbreaks
in southern California may have been introduced into
the United States from Mexico by importation of infected
domestic rabbits ( Vail and McKenney, 1943 ). The dis –
ease is endemic in the western United States, where the
Brush rabbit ( Sylvilagus bachmani ) is the natural reservoir
(Marshall and Regnery, 1960; Regnery and Miller, 1972 ).
Myxomatosis was introduced intentionally into
Australia in an effort to control what had become
Australia’s major animal pest, the European rabbit
(Oryctolagus cuniculus ). The virus was first introduced
into Australia in 1926 and used only in experimental
studies aimed at determining its feasibility as a control
measure for rabbits. In 1950, the virus was released into
the wild rabbit population where it decimated the rab –
bit population of the continent by 1953. The disease is
now endemic in the wild rabbit population of Australia,
where it occasionally assumes epidemic proportions
when climatic conditions favor vector activity. Within a
decade following release of Myxoma virus into the rabbit
population, it became evident that through natural selec –
tion genetically resistant strains of rabbits had emerged.
In these rabbits, a virulent strain of Myxoma virus caused
only 25% mortality compared to 90% mortality in non-
resistant strains of rabbits ( Fenner and Ratcliffe, 1965 ).
This selective pressure continues with regional increases
in resistance of Australian rabbits ( Williams et al., 1990 ).
Genetic modification of Myxoma virus was recog –
nized soon after its release into the rabbit population,
and by the fourth year markedly attenuated strains
of virus had replaced virulent virus as the dominant
strains. The naturally attenuated viruses caused a
milder disease of longer duration, which favored vector
transmission and thus persistence of the virus ( Fenner
and Woodroofe, 1965; Fenner et al., 1957 ). The evolution
of myxomatosis in Australia is a classic example of nat –
ural modification of both a virus and host until a state
of equilibrium is reached, permitting the continued
existence of both.
The introduction of myxomatosis into Europe fol –
lowed the early successes of the Australian campaign.
In 1952, while French officials were considering the
desirability of introducing the disease, a private indi –
vidual acquired the virus and released it on his own
estate in an effort to control the rabbit population. The

369 DNA VIRUS INFECTIONS
II. RABBITSvirus spread rapidly through the countryside, and by
the end of 1953 myxomatosis had been diagnosed in
Belgium, the Netherlands, Germany, Luxembourg,
Spain, and England ( Armour and Thompson, 1955;
Fenner and Ratcliffe, 1965; Lubke, 1968 ). Myxomatosis
is now endemic in rabbits of the genus Sylvilagus in
both North and South America and in wild Europen
rabbits ( Oryctolagus cuniculus ) in South America,
Europe, Australia, and New Zealand.
ETIOLOGY
Myxomatosis is caused by one of several strains of
the species, of which Myxoma virus is the type species
of the genus Leporipoxvirus in the Chordopoxvirinae sub –
family of the family Poxviridae (Fauquet et al., 2005 ).
Antigenic differences, demonstrated among different
strains of the virus ( Fenner, 1965; Reisner et al., 1963 ),
have prompted some to consider the California strains
of virus as distinct from Myxoma virus , and the desig –
nation “California rabbit fibroma virus” has been used
to describe this virus. However, the demonstrable
antigenic differences are insufficient to justify this dis –
tinction, and California strains of virus are considered
strains of Myxoma virus (Fauquet et al., 2005 ).
Myxoma virus is antigenically closely related to the
Rabbit fibroma virus as demonstrated by agar-gel dif –
fusion microprecipitation techniques ( Fenner, 1965 ).
Heat-inactivated Myxoma virus has been reactivated by
fibroma virus ( Berry and Dedrick, 1936; Fenner, 1962 ),
further demonstrating the close relationship between
these two viruses. The Berry-Dedrick phenomenon of
poxvirus reactivation was confirmed by Smith (1952) ,
who demonstrated a spectrum of virulence for strains
of myxoma and fibroma viruses. Fenner and Marshall
(1957) , in a study involving 92 strains of virus, estab –
lished a virulence spectrum ranging from strains causing
over 99% mortality in European rabbits to others caus –
ing less than 30% mortality. The most virulent strains
were the Standard Laboratory, Lausanne, and California
strains, whereas the least virulent were the neuromyx –
oma and Nottingham strains. Ecological pressures such
as those previously described in Australia could have
been responsible for the emergence of many of these
strains of viruses. In many instances, however, viruses
have been manipulated in the laboratory to the point of
permanent modification ( Kilham, 1957, 1958 ). The entire
genome of the Lausanne strain has been sequenced
(Cameron et al., 1999 ). Infection of Oryctolagus cuniculus
with Myxoma virus has provided a model system for elu –
cidating the immunopathogenesis of poxvirus in the host
(Stanford et al., 2007 ).
The chemical and physical characteristics of Myxoma
virus have been described (Andrewes and Porterfield,
1989; Fenner, 1953; Fenner and Ratcliffe, 1965 ). Myxoma
virus is readily propagated at 35°C on the chorioallantoic membrane of embryonated hens’ eggs, forming distinct
pocks ( Fenner and McIntyre, 1956 ). Different strains of
virus cause pocks of various sizes, the variation being
sufficiently distinct to allow tentative strain identifica –
tion. The South American strains cause large pocks,
whereas the California strains produce small focal
lesions on the membrane ( Fenner and Marshall, 1957 ).
The virus can also be propagated in cell cultures derived
from rabbits and other species, including chicken, squir –
rel, rat, hamster, guinea pig, and human (Andrewes and
Porterfield, 1989; Woodroofe and Fenner, 1965 ). Distinct
differences in plaque size on rabbit kidney cell cultures
can be demonstrated between the South American
and California strains, the former causing much larger
plaques ( Woodroofe and Fenner, 1965 ). The most sensi –
tive method for isolation of Myxoma virus under labo –
ratory conditions is inoculation of the skin of European
rabbits ( Fenner and McIntyre, 1956 ).
EPIDEMIOLOGY
Myxomatosis is endemic on four continents:
Australia, Europe, North America, and South America.
In Brazil and Uruguay, the virus is endemic in wild
rabbits ( Sylvilagus ), particularly Sylvilagus brasiliensis
(Aragao, 1943). A similar situation may exist in Panama
and Colombia, where the strains of virus are similar in
virulence to South American strains, but antigenically
more closely related to the California strains ( Fenner,
1965 ). In the forested area of Argentina, the virus is also
endemic in Sylvilagus rabbits, but in the southern part
of the country and in Chile the principal reservoir is the
wild European rabbit ( Oryctolagus cuniculus ) (Fenner
and Ratcliffe, 1965 ). The California strains of Myxoma
virus , also known as the California rabbit fibroma virus
(Andrewes and Porterfield, 1989), are endemic in wild
rabbits ( Sylvilagus ), especially Sylvilagus bachmani,
which serves as the principal source of infection for
domestic rabbits ( Marshall et al., 1963 ). A similar situ –
ation exists in Mexico, where Sylvilagus bachmani has
been shown to be the reservoir in Baja California ( Licón
Luna, 2000 ). In Australia, the Myxoma virus has been
endemic in wild European rabbits ( Oryctolagus cunicu –
lus) since its introduction into the rabbit population
in 1950. Following the introduction of the virus into
France in 1952, myxomatosis has become established
in most countries of Europe, the wild European rab –
bit serving as the predominant host species. In Europe,
Myxoma virus and Rabbit hemorrhagic disease virus occur
endemically in the same free-living populations of
Oryctolagus cuniculus (Calvete et al., 2002 ) and seroposi –
tivity to both viruses occurs significantly suggesting
the possibility of predisposition ( Marchandeau et al.,
2004 ). In rabbit farms in Belgium, the Netherlands,
and Germany, the seroprevalence of Myxoma virus was

II. RABBITS
14. VIRAL DISEASES
370
higher in herds with recurrent respiratory or digestive
diseases ( Marlier et al., 2001 ).
The naturally susceptible species are the European
rabbit ( Oryctolagus cuniculus ), the European hare ( Lepus
europaeus ), the Mountain hare ( Lepus timidus ), the
Forest rabbit ( Sylvilagus brasiliensis ), the Brush rabbit
(Sylvilagus bachmani ), and the Eastern Cottontail rab –
bit ( Sylvilagus floridanus ). Experimentally, several addi –
tional species of Sylvilagus can be infected ( Fenner and
Ratcliffe, 1965; Regnery and Marshall, 1971 ).
The principal mode of transmission of the virus is
mechanical transport of virus on mouth parts by arthro –
pod vectors, mosquitoes and fleas being most often
incriminated ( Grodhaus et al., 1963 ), but biting flies,
gnats, mites, and lice may serve as vectors ( Fenner and
Woodroofe, 1953; Mykytowycz, 1957 ). The source of
virus is usually the superficial layers of the skin, espe –
cially of the eyelids and at the base of the ears ( Fenner
and Woodroofe, 1953 ), where surface-feeding arthro –
pods obtain the virus and serve as mechanical vec –
tors. Experimentally, virus can spread by contact in the
absence of arthropod vectors, and such transmission
may occur under natural conditions in rabbit warrens
(Mykytowycz, 1958, 1961 ). An outbreak in Hungary at
a rabbit farm was attributed to airborne transmission
as the disease occurred in winter when mosquitoes and
fleas were uncommon ( Farsang et al., 2003 ). Windborne
spread was suspected in France ( Arthur and Louzis,
1988 ). Transmission of Myxoma virus by contaminated
spines of thistles ( Circium vulgare ) has been described
(Dyce, 1961; Mykytowycz, 1961 ). The claws of preda –
tory birds and carrion feeders, such as buzzards and
crows, may be contaminated with virus, and such birds
may play a role in dissemination of the virus ( Borg and
Bakos, 1963 ).
In an attempt to improve the usefulness of myxomato –
sis in rabbit control, the European rabbit flea, Spilopsyllus
cuniculi, was introduced into Australia in 1966 ( Sobey
and Menzies, 1969 ). The flea reproduced in wild rabbit
populations and transmitted both introduced and field
strains of Myxoma virus (Shepherd and Edmonds, 1977;
Sobey and Conolly, 1971 ). As a result of flea introduction,
myxomatosis has become more prevalent in drier table –
land areas ( Parer and Korn, 1989 ), and outbreaks of myx –
omatosis have shifted from summer to spring ( Shepherd
and Edmonds, 1978; Shepherd et al., 1978 ). In France,
there is evidence that mosquitoes of the genus Anopheles
are the principal vectors of summer epidemics. The rab –
bit flea, Spilopsyllus cuniculi, is probably a major vector,
especially during winter months when mosquito activity
is low ( Fenner and Ratcliffe, 1965 ).
Myxomatosis in Britain is characterized by milder
seasonal fluctuations in disease incidence than in
Australia, California, and France ( Ross and Tittensor,
1986 ). Mosquitoes play a minor role as vectors, whereas the rabbit flea, Spilopsyllus cuniculi, which is
less influenced by seasonal changes, is the major vector
(Andrewes et al., 1959; Armour and Thompson, 1955;
Lockley, 1954; Mead-Briggs, 1964 ). The Myxoma virus
in Britain has not undergone the rapid loss of virulence
observed with the Australian and French viruses. The
increase in resistance to myxomatosis in wild rabbit pop –
ulations has resulted in the appearance of more virulent
strains of Myxoma virus (Ross and Sanders, 1987 ). While
mildly virulent strains have emerged in Britain, the pre –
dominant strains are moderately virulent ( Chapple and
Bowen, 1963; Chapple and Lewis, 1964; Fenner and
Chapple, 1965 ), with recent estimates of between 47%
and 69% mortality in infected rabbits ( Ross et al., 1989 ).
The different evolution of Myxoma virus in Britain has
been attributed to the fact that the virus is predominantly
flea-transmitted. The flea is less seasonal and less mobile
than the mosquito. That fleas move in large numbers
from dead animals while moving only occasionally from
live ones would seem to favor transmission of virulent
virus strains ( Fenner and Marshall, 1957 ). The propor –
tion of infective fleas produced is inversely related to the
survival time of rabbits following infection ( Mead-Briggs
and Vaughan, 1975 ). The flea is also an effective reservoir
of virus, possessing a longer life span than mosquitoes.
The life span of active female mosquitoes is usually 2–3
weeks, whereas fleas have been known to feed actively
for over 1 year. The Myxoma virus can persist for 105 days
in rabbit fleas with no rabbit contact in artificial burrows
(Chapple and Lewis, 1965 ).
In experimental studies of viral pathogenesis and host
resistance, a virulent strain of Myxoma virus (SLS) inocu –
lated into laboratory European rabbits ( Oryctolagus cunic –
ulus) resulted in fatal infections whereas wild European
rabbits with naturally acquired innate resistance recov –
ered ( Best and Kerr, 2000 ). Conversely, inoculation of lab –
oratory European rabbits with an attenuated, naturally
derived field strain of Myxoma virus resulted in recovery
from infection and rabbits with innate resistance expe –
rienced only mild disease. A similar study examined
the pathogenesis of two Californian strains of Myxoma
virus es (MSW and MSD) in European rabbits and found
that both susceptible rabbits and rabbits that had natu –
rally acquired innate resistance, experienced acute fatal
infections with the MSW strain, whereas, the MSD strain
caused classical signs of myxomatosis in both strains of
rabbits ( Silvers et al., 2006 ).
CLINICAL SIGNS
Considerable differences in the virulence of Myxoma
virus strains complicate discussion of the clinical dis –
ease, as does the fact that different species and strains
of rabbits vary considerably in susceptibility to Myxoma
virus . Major emphasis is given to discussion of disease

371 DNA VIRUS INFECTIONS
II. RABBITSin Oryctolagus by the major strains of virus found in
Australia, California, Europe, and South America.
SIGNS IN SYLVILAGUS SPECIES Rabbits of the genus
Sylvilagus, the natural host of the virus, are relatively
resistant to infection and generally develop skin tumors
and lesions, but rarely systemic disease, except in young
rabbits (Aragao, 1943). Generally, the nodules occur at
the site of infection, persist for a variable amount of time,
and then regress. The size of the skin nodule and its per –
sistence vary with the strain of rabbit and the strain of
virus ( Fenner and Ratcliffe, 1965; Marshall and Regnery,
1963; Regnery and Miller, 1972; Silvers et al., 2009 ).
SIGNS IN LEPUS SPECIES The European hare (Lepus
europaeus) is resistant to Myxoma virus under experi –
mental conditions, and field experience supports this
observation. Occasionally, however, individual hares
(Lepus europaeus and Lepus timidus ) with mild to severe
generalized myxomatosis have been encountered
(Fenner and Ratcliffe, 1965 ).
SIGNS IN ORYCTOLAGUS CUNICULUS Myxoma
virus infection in the European rabbit usually results in
severe disease with high mortality ( Marshall et al., 1963;
Patton and Holmes, 1977 ). The severity and variety of
clinical disease is largely determined by the strain of
virus as well as the strain of rabbit ( Sobey, 1969; Sobey
et al., 1970 ) and have been described in detail ( Fenner
and Marshall, 1957; Fenner and Ratcliffe, 1965 ). The dis –
cussion which follows is a summation of the findings of
these workers and others ( Chapple and Bowen, 1963;
Kessel et al., 1931 ).
Signs which develop following infection with
California strains of the virus in susceptible rabbits can
be categorized as peracute, acute, or chronic. Rabbits
with the peracute form of disease die within 1 week
after exposure to the virus, exhibiting only edema of the
eyelids and lethargy prior to death. In the acute form of
disease, in which rabbits survive for 1–2 weeks, usually
edema of the eyelids, resulting in a “droopy” appear –
ance of the eyes, appears at 6–7 days. Inflammation
and edema around the anal, genital, oral, and nasal
orifices are also observed. Skin hemorrhages and con –
vulsions precede death on the ninth or tenth day. The
few rabbits which survive beyond 10 days and have a
chronic form may develop purulent blepharoconjuncti –
vitis and edema at the base of the ears, signs more often
associated with other myxoma strains ( Figure 14.1 ).
The nodule which develops at the site of inoculation is
not a clearly defined tumor, and under natural condi –
tions the development of myxomas is not characteris –
tic. Although nodules on the ears, head, and legs have
been reported ( Kessel et al., 1934 ), other workers have been unable to induce nodule development ( Fenner and
Marshall, 1957 ).
The original South American isolate of Moses (1911) ,
results in an acute disease with a mean survival time of
11 days. From 3–4 days following inoculation or natu –
ral infection with virus, a primary tumor may become
evident, and generalized tumors usually appear by the
sixth or seventh day. Edema of the eyelids occurs fol –
lowed by mucopurulent blepharoconjunctivitis, often
resulting in complete closure of the eyes. Mucopurulent
nasal discharge and pronounced edema of the base of
the ears, the perineal region, the external genitalia, and
lips are frequently seen. By the tenth day, hard convex
lumps may cover the body, head, and ears and occa –
sionally the legs. The lumps are not sharply demarcated
but may become markedly congested and ultimately
necrotic in rabbits surviving for 2 weeks. Dyspnea is
often seen in protracted cases, but appetite may be
maintained until shortly before death. Terminal convul –
sions frequently precede death, which usually occurs
8–15 days after infection. Infection with less virulent
South American and Australian strains results in milder
disease with less edema and nasal and ocular discharge,
more clearly demarcated nodules, and lower mortality.
The laboratory-attenuated neuromyxoma virus induces
a mild disease with little or no mortality ( Hurst, 1937b ).
The predominant myxoma strains in Europe are the
virulent Lausanne strain and its naturally attenuated
derivatives originating from the virus introduced into
France from Brazil in 1952. The more virulent European
viruses cause severe disease in rabbits, resulting in mor –
tality of up to 100%, but modified strains which have
emerged are of lower morbidity and mortality ( Arthur
and Louzis, 1988 ). With some of the naturally attenu –
ated British viruses, mortality is also decreased, and
tumors are flat rather than convex, resembling some of
the attenuated field strains in Australia ( Chapple and
Bowen, 1963 ). In France and Belgium, two forms of
the disease are recognized, nodular and amyxomatous
forms ( Marlier et al., 1999 ). The former is characterized
by florid skin lesions and severe immunosuppression,
accompanied by bacterial infections of the respiratory
tract, whereas in the latter, skin nodules are few and
small, with acute respiratory distress and copious nasal
discharge.
PATHOLOGY
The gross and microscopic pathology of myxoma –
tosis has been comprehensively reviewed ( Fenner and
Ratcliffe, 1965; Hurst, 1937a; Rivers, 1930 ). In adult
Sylvilagus, Myxoma virus usually causes localized skin
tumors. The tumors resemble the fibromas in European
rabbits produced by the rabbit fibroma virus (see later).
Hares or young Sylvilagus usually develop a mild
localized infection, although disseminated cutaneous

II. RABBITS
14. VIRAL DISEASES
372
FIGURE 14.1 Rabbit with myxomatosis displaying facial edema
with mucopurulent conjunctivitis. Courtesy of Dr. G. Van Hoosier.
FIGURE 14.2 Myxoma virus expressing lacZ-infected primary
skin lesion immunostained with antibody to β-galactosidase demon –
strates the wide distribution of the virus within the dermis and subcu –
tis. Hematoxylin and eosin counterstain. Courtesy of Dr. G. McFadden.fibromatous to myxomatous nodules similar to those
in acute myxomatosis may be found. Prominent gross
lesions in European rabbits with myxomatosis are
skin tumors (not characteristic of the California dis –
ease) and pronounced cutaneous and subcutaneous
edema, especially of the face and around body orifices.
Hemorrhages of the skin, heart, and subserosa of the
gastrointestinal tract may be observed, especially fol –
lowing infection with the California virus.
Lesions in the skin involve epithelial cells, fibroblasts,
and endothelial cells and range from proliferative to
degenerative, depending on the strain of virus. The skin
tumors result from proliferation of undifferentiated mes –
enchymal cells, which become the characteristic large
stellate (myxoma) cells surrounded by a homogeneous
matrix of mucinous material interspersed with capillar –
ies and inflammatory cells ( Figure 14.2 ). There is typi –
cally vascular endothelial proliferation and hypertrophy
with narrowing of the lumen ( Hurst, 1937a; Percy and
Barthold, 2007 ). Central necrosis of myxomas may be
attributed to occlusion of blood vessels by endothelial
proliferation. Epithelial cells overlying the tumor may
appear normal in early tumors, or show hyperplasia or
degeneration. Intracytoplasmic inclusions, in various
cells types, are especially prominent in the epidermis
(Patton and Holmes, 1977; Rivers and Ward, 1937 ).
Lesions in other organs, although not consistently
present, reflect the potential generalized nature of myx –
omatosis. Cellular proliferation, invariably present in
the skin, has also been described in pulmonary alveo –
lar epithelium and in reticulum cells of lymph nodes
and spleen ( Hurst, 1937a ). Focal hemorrhages may be
observed in skin, kidneys, lymph nodes, testes, heart,
stomach, and intestines. Degeneration and necrosis
occur frequently in lymph nodes, pulmonary alveoli,
spleen, and central veins of hepatic lobules. Stellate
cells may occur in lymph nodes, bone marrow, uterus,
ovaries, testes, and lungs ( Marcato and Simoni, 1977 ). In a survey of 66 rabbits, presumably Oryctolagus cunic –
ulus, that died of respiratory disease , Myxoma virus was
isolated from seven (10%), mainly from rabbits with
acute hemorrhagic pneumonia ( Marlier et al., 2000 ).
The mechanisms by which the virus manifests the
development of disease in rabbits have been described
(Stanford et al., 2007 ).
DIAGNOSIS
Myxomatosis in European rabbits can usually be
diagnosed by the clinicopathological features, particu –
larly the nodular form, due to the characteristic skin
lesions. Infection with the California viruses may be
harder to diagnose, however, owing to the frequent
absence of skin nodules and other signs of disease.
Diagnosis should be confirmed by virus isolation.
Intracutaneous inoculation of young susceptible rabbits
with fresh tissue collected from lesions free of bacterial
contamination results in lesions at the site of inoculation
within 1 week. The virus can be isolated by chorioal –
lantoic membrane inoculation of 11–13-day-old embry –
onated chicken eggs followed by incubation at 35°C
for 4–6 days. Distinct focal pocks develop if the virus
is present. The South American viruses cause large
pocks, the California virus intermediate-sized pocks,
and the fibroma virus minute pocks. Virus isolation on
chicken embryo fibroblast, Vero or RK13 cells can also
be accomplished. A cytopathic effect typical of poxvi –
ruses usually develops in 1–2 days but may take up to 7
days with some strains. The virus isolated can be iden –
tified as Myxoma virus by direct fluorescent antibody
test ( Takahashi et al., 1958, 1959 ), the plaque-neutral –
ization test ( Woodroofe and Fenner, 1965 ), the agar-gel
diffusion microprecipitation test ( Fenner, 1965 ), or the

373 DNA VIRUS INFECTIONS
II. RABBITSindirect immunoperoxidase test ( Marlier et al., 1999 ).
Myxoma virus DNA in tissue samples can be detected
by polymerase chain reaction (PCR) ( Barcena et al.,
2000; Kritas et al. 2008; Pérez de Rozas et al., 2008 ).
Rabbits that survive infection develop antibodies to
the virus, which can be detected by an enzyme-linked
immunosorbent assay (ELISA) that has been shown to
be more sensitive and specific than agar gel immuno –
diffusion, complement fixation (CF), or indirect immu –
nofluorescence ( Gelfi et al., 1999 ). Serum neutralization
has also been used to detect antibodies ( Marlier et al.,
1999 ). Infection of Sylvilagus rabbits with Myxoma virus
clinically resembles fibromatosis and should be differ –
entiated from the latter disease by PCR or inoculation
of young susceptible rabbits of the genus Oryctolagus.
Myxomatosis causes severe to fatal disease, whereas
fibromatosis causes a localized fibroma.
CONTROL
Control of myxomatosis is of prime importance in areas
where the virus is endemic in wild rabbit populations.
V ector control, including adequate screening to exclude
mosquitoes, serves to keep the disease under control.
Newly introduced rabbits should be quarantined in an
insect-proof facility for 2 weeks. To prevent spread within
a colony , all sick rabbits should be isolated. Fibroma virus
has been used as a live vaccine for myxomatosis, but
results have been variable ( Fenner and Ratcliffe, 1965 ). A
live attenuated myxomatosis vaccine (the MSD strain)
results in a mild reaction followed by immunity persist –
ing for 9 months ( McKercher and Saito, 1964 ). Jiran et al.
(1970) found this virus to be too virulent for use as a vac –
cine and further attenuated it by serial passage in rabbit
kidney cell cultures. The additionally modified virus, des –
ignated MSD/B, was safe and highly immunogenic.
However, efficacy of a vaccine has yet to be demon –
strated in commercial rabbits. In an outbreak of atypical
myxomatosis in Hungary, fibroma vaccine was not pro –
tective, whereas in experimental studies, a live myxo –
matosis vaccine protected rabbits ( Farsang et al., 2003 ).
An assessment of the Borgi strain vaccine of Myxoma
virus was reported during an outbreak of myxomato –
sis in Greece ( Kritas et al., 2008 ). Two-month-old vacci –
nated pregnant does were obtained by two commercial
rabbitries, only one of which had vaccinated resident
does. Within several weeks of their arrival, an explo –
sive outbreak of myxomatosis with high morbidity and
mortality occurred on both farms in both unvaccinated
and vaccinated rabbits, both imported and resident,
with the Lausanne strain of Myxoma virus . Since vacci –
nated rabbits were affected within 6 months of vaccina –
tion, the efficacy of the vaccine was questionable.
Field trials using various strains have been conducted
in free-living rabbit populations. A field trial of a recom –
binant vaccine, using a naturally attenuated myxoma field strain (6918) that expressed a Rabbit hemorrhagic dis –
ease virus VP60 protein, conducted on an island off the
coast of Spain, revealed that the vaccine induced anti –
bodies to both viruses in vaccinated rabbits without any
adverse side effects ( Torres et al., 2001 ). Furthermore,
the vaccine exhibited limited horizontal transmission,
either by direct contact or fleas, as uninoculated rabbits
also exhibited seroconversion. Although the observa –
tion period was brief (32 days), the vaccine appeared to
be both safe and efficacious. An attenuated strain (SG33)
was used in a field trail in juvenile rabbits in France and
vaccinated rabbits had a 1.8-fold greater odds of surviv –
ing than unvaccinated rabbits over 4 years of surveil –
lance ( Guitton et al., 2008 ). In a field trial in Zaragoza
province in northeast Spain, young European rabbits
(Oryctolagus cuniculus ) born in the current year and
unvaccinated, were 13.6 times more likely to die than
rabbits vaccinated against both myxomatosis (POX-LAP)
and Rabbit hemorrhagic disease (CYLAP-VHD), over
a 90-day observation period after vaccination ( Calvete
et al., 2004 ).
Rabbit (Shope) Fibroma Virus
HISTORY
The transmissible tumor-inducing agent now known
as rabbit or Shope fibroma virus was isolated from an
Eastern Cottontail rabbit (Sylvilagus floridanus) in 1932
(Shope, 1932a ). The virus was transmissible to Eastern
Cottontail and European rabbits (Oryctolagus cuniculus),
producing localized fibromas in both species. Shope
(1932a) described the gross and microscopic lesions of
both the natural and experimental disease and identi –
fied the causative agent as a virus antigenically related to
Myxoma virus (Shope, 1932b ). Similarities between Rabbit
fibroma and Myxoma virus es have subsequently been
confirmed by cross-immunity tests ( Shope, 1936 ), ether
sensitivity ( Andrewes and Horstmann, 1949; Fenner,
1953 ), virus-reactivation studies ( Berry and Dedrick,
1936 ), microprecipitation procedures ( Fenner, 1965 ), and
sequence comparison ( Willer et al., 1999 ). The Rabbit
fibroma virus , initially believed to cause only localized
benign fibromas, was later shown to cause severe gen –
eralized disease in newborn European ( Duran-Reynals,
1940; Joiner et al., 1971 ) and Eastern Cottontail rabbits
(Yuill and Hanson, 1964 ). The disease is historically con –
sidered a benign endemic disease of wild Sylvilagus rab-
bits, of little economic significance to commercial rabbit
producers or laboratory investigators. However, an out –
break of fibromatosis in a commercial rabbitry resulting
in high morbidity and mortality in newborn rabbits was
reported in 1971 ( Joiner et al., 1971 ). Thus, the disease
can be a threat to commercial rabbits in areas where it
is endemic in wild rabbit populations and outdoor hus –
bandry practices permit contact with arthropod vectors.

II. RABBITS
14. VIRAL DISEASES
374
Another tumor-producing virus, designated malig –
nant rabbit fibroma virus, has been recovered from
Oryctolagus cuniculus with experimentally induced Shope
fibromas ( Strayer et al., 1983a ). Although the virus was
present in a stock of Rabbit fibroma virus , its origin is
unclear. The virus is antigenically similar to both Rabbit
fibroma and Myxoma virus es and is considered a recom –
binant of the two viruses ( Block et al., 1985 ). The virus
induces a syndrome of severe immunosuppression result –
ing in disseminated malignancy and opportunistic infec –
tions ( Corbeil et al., 1983; Skaletsky et al., 1984; Strayer
and Sell, 1983; Strayer et al., 1983a, 1983b, 1983c ). The
disease may be a useful model for the study of virus-
induced immunologic impairment ( Strayer et al., 1985 ).
ETIOLOGY
Rabbit fibroma virus is a Leporipoxvirus of the fam –
ily Poxviridae (Fauquet et al., 2005 ) and is closely related
to Myxoma virus (Fenner, 1953 ) and the hare and squir –
rel fibroma viruses ( Fenner, 1965 ). The complete genome
of Rabbit fibroma virus has been sequenced ( Willer
et al., 1999 ). The chemical and physical characteristics of
fibroma virus have been summarized by Gross (1983) .
The virus can be propagated on the chorioallantoic mem –
brane of chicken embryos, but characteristic lesions as
observed with Myxoma virus are not produced ( Gross,
1983 ). The virus has been propagated in cell cultures
derived from rats, guinea pigs, and humans ( Chaproniere
and Andrewes, 1957 ) and also in rabbit cell cultures
derived from Eastern Cottontail and domestic rabbits
(Constantin et al., 1956; Hinze and Walker, 1964; Kasza,
1974; Kilham, 1956; Padgett et al., 1962 ). Fibroma virus in
cultured rabbit kidney cells induces pronounced changes
in cell growth and morphology , and inoculation of virus-
transformed cells into the cheek pouch of hamsters
results in tumor formation ( Hinze and Walker, 1964 ).
EPIDEMIOLOGY
Since Rabbit fibroma virus was first isolated from an
Eastern Cottontail rabbit in New Jersey ( Shope, 1932a ),
the disease has been recognized in several other states,
as well as in Canada. Herman et al. (1956) found that
more than 50% of wild Sylvilagus rabbits trapped in the
Patuxent Wildlife Refuge in Maryland had Rabbit fibroma
virus or antibodies to Rabbit fibroma virus . The virus has
also been isolated from Sylvilagus rabbits in Wisconsin
(Yuill and Hanson, 1964 ), Michigan ( Herman et al., 1956 ),
and Ohio ( Kasza, 1974 ). Recognition of the disease in
Texas ( Joiner et al., 1971 ) indicates that it may be more
widespread in the U.S. than was formerly believed.
The natural transmission cycle of Rabbit fibroma virus
has not been completely elucidated. The virus may
persist in the epidermis of experimentally infected
Sylvilagus rabbits for 5–10 months ( Kilham and Dalmat, 1955; Kilham and Fisher, 1954 ), which would enhance
the likelihood of mechanical arthropod transmission.
Experimentally, several species of mosquitoes as well
as triatomes, fleas, and bedbugs can serve as vectors
(Dalmat, 1959; Kilham and Dalmat, 1955; Kilham and
Woke, 1953 ). Infected mosquitoes are capable of infect –
ing wild Sylvilagus rabbits ( Kilham and Dalmat, 1955 ).
Experimental and circumstantial evidence suggests that
the principal mode of transmission for fibroma virus is
biting arthropods, a situation similar to that described
for myxomatosis. Infection may also occur by direct
contact with skin injuries being a predisposing factor
(Grilli et al., 2003 ).
The natural host of fibroma virus is the Eastern
Cottontail rabbit ( Sylvilagus floridanus ). Three other spe –
cies of Sylvilagus (Sylvilagus bachmani, Sylvilagus nuttalli,
and Sylvilagus auduboni ) are refractory to fibroma virus
(Fenner and Ratcliffe, 1965 ). A closely related virus,
isolated from fibromas in Sylvilagus bachmani, failed
to induce lesions in other Sylvilagus species ( Marshall
and Regnery, 1960 ). This virus, considered a strain of
Myxoma virus , is a related virus named California Rabbit
fibroma virus (Andrewes and Porterfield, 1989). The
European rabbit ( Oryctolagus cuniculus ) is susceptible to
fibroma virus ( Shope, 1932a ) as is the Snowshoe hare,
Lepus americanus (Yuill, 1981 ), but the European hare,
Lepus europaeus, is refractory to the virus ( Fenner and
Ratcliffe, 1965 ). Whereas the European rabbit is readily
infected, it does not serve as a good source of infection
for mosquitoes because of low virus concentrations in
the skin ( Fenner and Ratcliffe, 1965 ).
CLINICAL SIGNS
The clinical signs are largely those described by
Shope (1932a, 1932b, 1936) . The tumors observed in
natural fibromatosis of Eastern Cottontail rabbits are
almost invariably on the legs or feet, usually one to
three tumors occurring on an infected rabbit. Tumors
may occur on the muzzle and around the eyes, and
measure up to 7 cm in diameter and 1–2 cm in thickness.
The tumors are subcutaneous, unattached to underly –
ing tissues, and may persist for several months, and in
some instances for nearly a year ( Kilham and Dalmat,
1955 ). Other clinical signs are usually absent, the tumor-
bearing rabbit remaining apparently normal through –
out the disease. Metastases do not occur ( Shope, 1932a ).
The clinical signs in experimentally infected European
rabbits resemble those in Sylvilagus rabbits. However,
regression of tumors is usually more rapid than in
Sylvilagus rabbits. Inoculation of newborn European
rabbits frequently results in fatal systemic infections
(Duran-Reynals, 1945 ). Systemic infections of adult
European rabbits have also been described ( Hurst,
1937c ), but the usual result of infection is the develop –
ment of localized benign tumors. In the only reported

375 DNA VIRUS INFECTIONS
II. RABBITSnatural outbreak of fibromatosis of European rabbits,
diffuse subcutaneous induration involving the underly –
ing tissues occurred ( Joiner et al., 1971 ). Subcutaneous
tumors were the predominant sign in adult rabbits but
suckling rabbits had lethargy and loss of condition, in
addition to the skin lesions. Hyperemia and edema of
the external genitalia of male and female rabbits were
also observed.
A survey of tumors in pet rabbits submitted to
the surgical biopsy service of a veterinary school in
Pennsylvania over a 16-year period revealed that Shope
fibroma was diagnosed in 19 tumors from 179 rabbits
(von Bomhard et al., 2007 ). The species of rabbits was
not provided and diagnosis was based on histopatho –
logic findings. The limbs, head, and pinna were affected
in 75% of cases. Case reports of keratitis with a peri-
limbal mass due to fibroma virus have been diagnosed
in individual European rabbits ( Oryctolagus cuniculus )
(Keller et al., 2007; McLeod and Langlinais, 1981 ).
PATHOLOGY
The gross and microscopic changes associated with
fibroma virus infection in naturally and experimentally
infected Sylvilagus rabbits and in experimentally infected
European rabbits have been described by several work –
ers ( Ahlstrom, 1938; Andrewes, 1936; Dalmat and
Stanton, 1959; Fisher, 1953; Grilli et al., 2003; Kilham and
Fisher, 1954; Shope, 1932a; Yuill and Hanson, 1964 ). The
pathological changes in the only reported natural out –
break of fibromatosis in European rabbits were described
by Joiner et al. (1971) . The earliest gross lesion in experi –
mentally infected Sylvilagus rabbits is a slight thickening
of the subcutaneous tissue, followed by development of
clearly demarcated soft swellings usually evident 6 days
after inoculation. The tumors enlarge, increase in density,
and usually reach maximum size by 12 days. The aver –
age size of the tumors is 4–6 cm with a thickness of 2 cm.
Tumors may persist for months before regressing, leav –
ing the rabbit essentially normal. Experimentally infected
newborn Sylvilagus rabbits may die of generalized fibro –
matosis. Under natural conditions, however, this form
of the disease has not been observed. Gross lesions in
experimentally infected European rabbits are similar to
those observed in Sylvilagus rabbits but regression of the
tumors occurs more rapidly.
The earliest microscopic lesions in Sylvilagus rab –
bits are an acute inflammation followed by localized
fibroblastic proliferation, accompanied by both mono –
nuclear and polymorphonuclear leukocyte infiltrations.
Fibroblasts proliferate until a distinct tumor is formed,
consisting of spindle-shaped and polygonal connective
tissue cells with abundant cytoplasm. Mitotic figures
are few. Many tumor cells may have large intracyto –
plasmic inclusions characteristic of poxvirus infections.
Mononuclear leukocyte cuffing of vessels adjacent to the tumor may be observed, and a pronounced accu –
mulation of lymphocytes at the base of the tumor is
often seen. Degeneration of the overlying epidermis
may result from pressure ischemia, followed by necro –
sis and sloughing of the epithelium and tumor. In many
instances, however, the tumors regress without epithe –
lial sloughing. Regression is usually complete within 2
months after appearance of tumors. Andrewes (1936)
described a strain of fibroma virus which caused a more
inflammatory and less proliferative lesion than the
fibroma virus isolated by Shope (1932a) .
Histologically, typical fibromas are observed in
European rabbits experimentally infected with fibroma
virus. The lesions differ only slightly from those in
Sylvilagus rabbits, the principal difference being the
absence of epidermal degeneration often observed in
the latter species ( Shope, 1932a ). However, Ahlstrom
(1938) described epidermal degeneration in European
rabbits. Microscopic lesions in the natural outbreak of
fibromatosis in European rabbits ranged from tumors
resembling myxomas to typical fibromas ( Joiner et al.,
1971 ). Eosinophilic cytoplasmic inclusions were occa –
sionally observed in tumor cells, and similar inclusions,
possibly of viral origin, were seen in epithelial cells
overlying the tumors. The lesions encountered in adult
and newborn rabbits did not differ significantly.
DIAGNOSIS
Fibromatosis should be differentiated from papilloma –
tosis and myxomatosis. Clinically, differentiation from
papillomatosis can be readily accomplished as fibromas
are essentially flat, subcutaneous, loose, rubbery tumors,
whereas papillomas are tumors of the skin which are
heavily keratinized and project in irregular fashion from
the skin surface. Histopathological differentiation is also
easily accomplished, especially when intracytoplasmic
inclusion bodies are observed. Historically, subcutane –
ous inoculation of young European rabbits with a tumor
cell suspension was used to differentiate the two viruses.
Rabbit fibroma virus induces local fibromas, whereas
Myxoma virus causes severe systemic and often fatal dis –
ease. Virus isolation in cell cultures or on the chorioallan –
toic membranes of chicken embryos should be attempted
to confirm the diagnosis. The virus can be identified by
serum neutralization tests ( Yuill, 1981 ).
CONTROL
Because the disease is endemic and of little signifi –
cance in Sylvilagus rabbits, no control measures have
been developed. Fibromatosis is also not an important
problem in domestic rabbits; however, since the disease
has been encountered in a rabbitry in an area where
the disease is apparently endemic in wild rabbits, con –
trol measures might be considered. In such areas, the

II. RABBITS
14. VIRAL DISEASES
376
method of choice for preventing infection of rabbits
held in outdoor enclosures is vector control. Careful
analysis of the role of possible vectors in natural out –
breaks should be made in preparation for the establish –
ment of vector control methods.
Hare Fibroma Virus
HISTORY
Epidemics of a fibromatous disease in European hares
(Lepus europaeus ) occurred in France and Northern Italy
in 1959 ( Lafenetre et al., 1960; Leinati et al., 1959 ). The
causative agent was a poxvirus related to Myxoma virus
(Leinati et al., 1961 ). In retrospect, a nodular skin disease
of hares in Germany designated hare sarcoma was prob –
ably hare fibromatosis ( Dungern and Coca, 1909 ).
ETIOLOGY
The causative agent of hare fibromatosis is a
Leporipoxvirus in the family Poxviridae (Fauquet et al.,
2005 ), which has been shown by plaque-neutralization
and cross-protection tests to be antigenically related to
myxoma, rabbit fibroma, and squirrel fibroma viruses
(Woodroofe and Fenner, 1962 ). Agar-gel diffusion micro –
precipitation techniques reveal that hare fibroma virus
shares more common antigens with Rabbit fibroma virus
than with Myxoma virus (Fenner, 1965 ). A considerable
degree of cross-protection exists between hare fibroma
and myxoma viruses. European rabbits immune to
Myxoma virus are completely refractory to hare fibroma
virus, whereas rabbits immunized with hare fibroma
virus develop signs when infected with Myxoma virus
but survive, indicating some protection ( Woodroofe and
Fenner, 1965 ).
EPIDEMIOLOGY
Hare fibromatosis has been recognized only in
Europe, where under natural conditions it infects the
European hare. However, dermal tumors were reported
on Cape hares ( Lepus capensis ) in the Laikipia District of
Kenya that had gross and histopathologic similarities to
hare fibromatosis ( Karstad et al., 1977 ). In 2001, an out –
break of fibromatosis occurred in farmed game European
hares ( Lepus europaeus ) in Italy ( Grilli et al., 2003 ). The
European rabbit ( Oryctolagus cuniculus ) is susceptible to
the virus, but natural outbreaks of disease in rabbits have
not been reported. A seasonal occurrence of disease has
been reported, with the peak incidence in late summer
and autumn ( Lafenetre et al., 1960; Leinati et al., 1959 ).
The mode of transmission and interepidemic survival of
the virus are unknown.
CLINICAL SIGNS
In European hares, the disease is characterized by
development of numerous skin nodules, up to 2.5 cm in size, occurring especially on the face, eyelids, and
around the ears. In farmed game European hares, fibro –
mas also occurred on the legs ( Grilli et al., 2003 ). The
nodules closely resemble rabbit fibromas ( Leinati et al.,
1961 ). In adult European rabbits the virus causes for –
mation of small fibromas, but newborn rabbits exhibit
large fibromas resembling the lesions of rabbit fibroma –
tosis ( Fenner and Ratcliffe, 1965 ).
PATHOLOGY
The gross and microscopic appearance of the lesions
of hare fibroma is similar to that of the lesions of rabbit
fibroma ( Lafenetre et al., 1960; Leinati et al., 1961 ).
DIAGNOSIS
The disease is usually diagnosed from clinicopatho –
logical features. The diagnosis can be confirmed by
virus isolation in rabbit kidney cell cultures or on
the chorioallantoic membrane of chicken embryos.
Serological characterization of the virus can be achieved
using the agar-gel diffusion technique ( Fenner, 1965 ).
CONTROL
Because the disease is endemic and of little signifi –
cance in hares, no control measures have been devel –
oped. Farmed game hares, used for stocking areas,
should be free of infection to avoid dissemination of the
virus ( Grilli et al., 2003 ).
Rabbitpox
HISTORY
Rabbitpox was first diagnosed at the Rockefeller
Institute in New York when a highly fatal disease
occurred spontaneously in the European rabbit
(Oryctolagus cuniculus ) colony in 1932 ( Greene, 1933,
1934a; Pearce et al., 1933; Rosahn and Hu, 1935 ). A
smaller outbreak had occurred in 1930. Rabbits devel –
oped an erythematous rash followed by cutaneous
eruptions closely resembling the pocks seen in human
infection with variola virus (smallpox). The disease was
extremely contagious and caused high mortality, espe –
cially in young rabbits and pregnant females. Belgian
hares were also susceptible to the disease. The causative
agent of the disease was a poxvirus ( Pearce et al., 1936;
Rosahn et al., 1936a ).
A spontaneous outbreak of a similar disease in a
laboratory rabbit colony in Holland was described by
Jansen (1941) . The disease was highly fatal and dif –
fered from the Rockefeller outbreak in that it was not
exanthematous, giving rise to the name “pockless”
rabbitpox ( Jansen, 1947 ). This disease was also caused
by a poxvirus closely related to vaccinia virus ( Jansen,
1946 ). A second outbreak of the disease in Holland was
reported by Verlinde and Wersinck (1951) . The second

377 DNA VIRUS INFECTIONS
II. RABBITSoutbreak of rabbitpox reported in the United States
occurred in New York ( Christensen et al., 1967 ) and
was also of the pockless type first observed by Jansen
(1947) in Holland. Another serious outbreak of the dis –
ease, resulting in mortality of 95%, occurred following
the introduction of supposedly inactivated rabbitpox
virus of the Dutch (Utrecht) strain into the rabbit colony
of a medical school ( Christensen et al., 1967 ). Various
aspects of the infection have been reviewed by Fenner
et al. (1989) . Rabbitpox virus infection in rabbits can
serve as a model for human smallpox infection as a fatal
systemic disease developed with a low inoculum and
inoculated rabbits naturally transmitted the virus to
susceptible rabbits by aerosol ( Adams et al., 2007 ).
ETIOLOGY
Rabbitpox virus is an Orthopoxvirus in the family
Poxviridae (Fauquet et al., 2005 ) and is antigenically
related to vaccinia virus ( Fenner, 1958; Hu et al., 1936;
Jansen, 1946 ). The biological properties of the Utrecht
(Jansen, 1941 ) and Rockefeller ( Greene, 1934a ) strains
of rabbitpox viruses are indistinguishable from certain
neurovaccinia strains ( Fenner; 1958; Fenner et al., 1989 ).
The close antigenic relationship between rabbitpox
virus and vaccinia virus, taken together with the fact
that all reported outbreaks of rabbitpox have occurred
in laboratory colonies, suggests that rabbitpox may be
a laboratory variant of vaccinia virus ( Greene, 1935a;
Verlinde and Wersinck, 1951 ). Wittek et al. (1977) used
genome mapping to show that rabbitpox virus (Utrecht
strain) was a strain of vaccinia virus. The viruses have
been shown to exhibit over 95% sequence similarity ( Li
et al., 2005 ).
Rabbitpox virus can be propagated on the chorioal –
lantoic membrane of chicken embryos with development
of distinct pocks. The predominant pock type is hemor –
rhagic ( Jansen, 1946 ), but white pock mutants have been
described ( Fenner and Sambrook, 1966; Fenner et al.,
1989 ). Rabbitpox virus has been propagated in several
cell lines, including HeLa, Chang Liver, L929 (mouse
fibroblast), human heart, KB (human epithelial), FL
(human amnion), AT (Chinese hamster epithelial), PK-2A
(pig kidney), and FAF cells (Chinese hamster fibroblast)
(Christensen et al., 1967 ). The Rockefeller strain of rab –
bitpox virus hemagglutinates chicken erythrocytes,
but the Utrecht strain and the strain isolated from the
first American outbreak of pockless rabbitpox do not
(Christensen et al., 1967 ).
EPIDEMIOLOGY
Rabbitpox is relatively rare, and all reported outbreaks
have occurred in laboratory colonies in the United States
and Holland. The highest mortality occurs in rabbits and
pregnant or lactating females ( Greene, 1934a, 1935b ).
Differences in susceptibility by rabbit breed also occur (Greene, 1935b ). Within infected colonies, the spread of
disease is extremely rapid, and, in the outbreak of 1932,
removal and isolation of infected rabbits failed to pre –
vent the disease from spreading throughout the colony
(Greene, 1934a ). The virus appears to spread by nasal
discharges, which usually appear on the third day after
infection. Airborne droplets may be inhaled or ingested
by susceptible rabbits ( Bedson and Duckworth, 1963 ).
Recovery from infection does not appear to result in
establishment of a carrier state, as recovered rabbits
can be safely mated with susceptible rabbits and clean
colonies derived from recovered stock ( Greene, 1934a ).
Arthropods have not been shown to play a role in the
epidemiology of rabbitpox infection. In general, the pri –
mary sources of infection resulting in outbreaks of dis –
ease have not been determined, although the origin may
have been rabbits inoculated with vaccinia virus ( Greene,
1935a ).
CLINICAL SIGNS
The clinical disease has been described in detail
(Bedson and Duckworth, 1963; Christensen et al., 1967;
Greene, 1934a ). The virus initially infects and multi –
plies in the nasal mucosa and later in lymph nodes of
the respiratory tract and in the lungs and spleen. Fever
and a profuse nasal discharge usually occur 2–3 days
after infection. Another early sign is enlargement and
induration of the lymph nodes, especially the popliteal
and inguinal nodes, which usually persist throughout
the course of disease. Widely distributed skin lesions
usually appear about 5 days after infection, initially as
an erythematous macular rash which becomes papular
and may progress to nodules up to 1 cm in size. These
nodules eventually form dry, superficial crusts. Macules
and papules may also occur on the mucous membranes
of the oral and nasal cavities. Extensive edema of the
face and oral cavity is often observed, as is focal necro –
sis of the hard palate and the gums. Hemorrhages in the
skin may occur in severe cases.
Male rabbits frequently develop severe orchitis with
extensive scrotal edema, and papules in the prepuce and
urethra are observed. Similar lesions may also occur in
the vulvae of females. If edema is extensive, urine reten –
tion may occur in either sex. Pregnant females usually
abort. The eyes are almost invariably affected and exhibit
blepharitis, purulent conjunctivitis, and acute keratitis
with corneal ulceration. Death usually occurs 7–10 days
after infection but may occur as early as 5 days, or rab –
bits may survive for several weeks before dying.
The generalized disease syndrome described above
represents the findings of Greene (1934a) and Bedson
and Duckworth (1963) with the Rockefeller strain of
virus. This strain may occasionally result in peracute
disease in which death is preceded only by fever,
anorexia, and occasionally blepharitis. The peracute

II. RABBITS
14. VIRAL DISEASES
378
form is unusual with the Rockefeller strain, but natu –
ral outbreaks of rabbitpox of the pockless type ( Jansen,
1941, 1946, 1947; Christensen et al., 1967 ), without ery –
thema or skin lesions, have occurred . In the first Dutch
outbreak, some rabbits died within 1 week after infec –
tion, with only anorexia, fever, and lethargy ( Jansen,
1941 ). In the first American outbreak of pockless dis –
ease, rabbits developed conjunctivitis and diarrhea and
died 7–9 days after experimental infection ( Christensen
et al., 1967 ). Jansen (1941, 1946, 1947) and Christensen
et al. (1967) described the occasional presence of scat –
tered papules on the lips and tongues of rabbits with
pockless rabbitpox. Experimentally, both the Utrecht
and Rockefeller strains of virus produce skin lesions
(Bedson and Duckworth, 1963 ).
PATHOLOGY
The gross and microscopic pathology of rabbitpox
has been described in detail by Greene (1934b) for the
Rockefeller Institute outbreak and by Christensen
et al. (1967) for the first outbreak of pockless rabbitpox
in the United States. The most distinctive gross lesions
are the skin lesions, which may range in severity from
a few localized papules to severe, almost confluent
skin lesions with extensive necrosis and hemorrhage.
Nodules occur in the mouth, upper respiratory tract,
spleen, liver, and lungs but may also occur elsewhere
in the body. Subcutaneous and oral submucosal edema
and edema of other body orifices are common. Only
rabbits with severe lesions of the mouth are emaciated
at necropsy.
The liver is enlarged, yellowish, and has numerous
small, gray nodules. Focal areas of hepatic necrosis may
be seen. Small nodules may also occur in the gallbladder.
The spleen is usually moderately enlarged with occa –
sional focal nodules or areas of focal necrosis. Scattered
diffusely throughout the lungs may be small white nod –
ules and, in advanced cases, focal areas of necrosis. The
testicles, ovaries, and uterus frequently exhibit diffuse
white nodules and marked edema. Necrosis of the tes –
tes occurs frequently, and the uterus may contain focal
abscesses. Focal nodules may be present in lymph nodes,
adrenal glands, thyroid glands, parathyroid glands, peri –
toneum, omentum, and, rarely, the heart. Specific gross
lesions are seldom observed in the central nervous sys –
tem or kidneys. In the pockless form of the disease, a few
pocks may be found in the mouth, and occasional skin
lesions may be revealed by shaving the rabbit. The prom –
inent gross lesions at necropsy are pleuritis, multifocal
hepatic necrosis, splenomegaly, and edema and hemor –
rhage of the testes. The small white nodules, abundant in
the more typical form of the disease, occur occasionally
in the lungs and adrenal glands.
The predominant histological lesion in rabbitpox
is the papule or nodule which occurs in the skin and many other organs. A typical nodule consists of a cen –
tral zone of necrosis, surrounded by mononuclear cells.
Adjacent tissues are frequently edematous and occa –
sionally hemorrhagic. Diffuse lesions with massive
mononuclear cell infiltration, necrosis, hemorrhage,
and edema often occur. Typical variola-like vesicles and
pustules are not characteristic of rabbitpox infection.
Vascular occlusion from the pronounced swelling of the
endothelium may result in necrotic lesions.
In the lungs, focal nodular lesions and diffuse pneu –
monitis, with perivascular mononuclear and poly –
morphonuclear cell infiltration occur. Focal to diffuse
pulmonary necrosis may be found. There is severe
congestion of the spleen, with marked distention of
sinuses by mononuclear cells, edema of Malpighian
corpuscles, and focal to diffuse necrosis. Lymph
nodes are generally greatly enlarged, mainly from
severe edema. Extensive necrosis of lymph nodes and
other lymphoid tissues such as Peyer’s patches may
occur. Hemorrhage and necrosis of the bone marrow
occur frequently, interspersed with areas of mono –
nuclear cell infiltration. Degeneration and necrosis of
hepatic parenchyma may be focal or diffuse and may
involve the whole organ. Focal necrosis with edema is
detected in the testes, as are necrotic foci in the adrenal
glands, uterus, thyroid glands, thymus, and salivary
glands. The characteristic cytoplasmic inclusions asso –
ciated with poxvirus infections are seldom encoun –
tered in rabbitpox.
DIAGNOSIS
Rabbitpox can be diagnosed by the clinical signs
and the characteristic gross and microscopic lesions.
Confirmatory diagnosis can be made by detection of
viral antigen in tissues by fluorescent antibody on tis –
sue impression smears ( Christensen et al., 1967 ) or by
virus isolation and identification. A PCR assay, using
four primers, followed by electrospray ionization mass
spectrometry, has been described which identified and
speciated orthopoxviruses ( Eshoo et al., 2009 ). The virus
can be isolated by chorioallantoic membrane inocula –
tion of chicken embryos or by cell culture propagation
of the virus on cells derived from rabbits, mice, or any
of several animal species ( Christensen et al., 1967 ). The
virus may be identified by the fluorescent antibody pro –
cedure, hemagglutination inhibition (some strains), or
cross-protection tests using vaccinia-immunized and
susceptible rabbits. Vaccinia-immunized rabbits should
be resistant, whereas severe disease with high mortality
should occur in susceptible rabbits.
CONTROL
Because the natural source of virus causing out –
breaks has not been determined, control measures
to prevent the occurrence of disease have not been

379 DNA VIRUS INFECTIONS
II. RABBITSdeveloped. In outbreaks, preventing spread of the
virus in the colony by isolation of sick rabbits has met
with mixed success ( Christensen et al., 1967; Greene,
1934a ). Investigators using rabbitpox virus experimen –
tally should take exceptional precautions to prevent
the virus from reaching susceptible rabbit populations.
In an outbreak in a large colony, vaccination with vac –
cinia virus can be used to protect uninfected rabbits
(Appleyard and Westwood, 1964; Boulter et al., 1971;
Rosahn et al., 1936b ).
Herpesvirus Infections
Herpesviruses have long been recognized as the
causative agents of respiratory and genital diseases
in humans, cattle, horses, and swine. They are also
the recognized causes of other disease syndromes in
many species of animals, including neoplastic diseases
in frogs, chickens, rabbits, monkeys, and humans. An
important characteristic of the herpesviruses is the
capacity to cause mild or subclinical disease after which
viral persistence in a latent state may ensue. Stresses of
various kinds may result in viral recrudescence even
after prolonged periods of latency. The possible exis –
tence of latent virus infections may have an important
influence on the quality of experimental animals and
potentially on the validity of experimental findings. In
addition, there are numerous examples of herpesviruses
that cause severe disease when infecting an abnormal
host as demonstrated when Human herpesvirus 1 infects
rabbits. The Herpesviridae of rabbits and hares can be sub –
divided into Gammaherpesvirinae and Alphaherpesvirinae
(Fauquet et al., 2005 ). The Gammaherpesvirinae con –
tain two tentative members of the genus Rhadinovirus ,
Leporid Herpesvirus 1 and Leporid Herpesvirus 2. Two
members of the Alphaherpesvirinae , genus Simplex virus ,
are known to infect rabbits, Leporid Herpesvirus 4 and
Human herpesvirus 1 .
Leporid Herpesvirus 1
HISTORY
Leporid herpesvirus I (synonyms Herpesvirus syl –
vilagus, cottontail virus, Hinze herpesvirus lymphoma)
was isolated from primary kidney cell cultures from
apparently healthy weanling Eastern Cottontail rabbits
(Sylvilagus floridanus ) trapped in Wisconsin ( Hinze, 1968,
1971a ). The virus was detected when focal areas of cell
destruction were observed in cell monolayers 14 days
after incubation. The virus has since been propagated in
kidney cells of both Eastern Cottontail and domestic rab –
bits. Another herpesvirus, apparently distinct from the
original isolate of Hinze (1971a) , was recovered from pri –
mary kidney cell cultures of Sylvilagus floridanus (Cebrian
et al., 1989 ).ETIOLOGY
The virus possessed the physical, chemical, and bio –
logical properties of a herpesvirus and was named
Herpesvirus sylvilagus (Heine and Hinze, 1972; Hinze,
1971a ). Although infectious virus could not be demon –
strated in lymphocytes from infected Sylvilagus rabbits,
it could be detected after in vitro cocultivation with rab –
bit kidney cells ( Hinze and Wegner, 1973; Wegner and
Hinze, 1974 ). The virus is strongly cell-associated ( Ley
and Burger, 1970 ), and both B and T lymphocytes are
infected ( Kramp et al., 1985 ). The virus can be propa –
gated in cells of both Eastern Cottontail and domestic
New Zealand White rabbits but not in cells from humans,
monkeys, hamsters, mice, nor in chicken embryos. The
highest concentrations of virus are obtained by the
use of primary kidney cell lines established from new –
born or juvenile Eastern Cottontail rabbits ( Cohrs and
Rouhandeh, 1982; Medveczky et al., 1984 ). The virus pos –
sesses no antigenic relationship to Leporid herpesvirus
2 or to four other Leporid herpesviruses ( Hinze, 1971a ).
The genome of the virus is similar to those of Herpesvirus
saimiri and Epstein-Barr virus, in that it contains a vari –
able number of repetitive DNA elements at both ends
(Heine and Hinze, 1972; Cohrs and Rouhandeh, 1987;
Rouhandeh and Cohrs, 1987; Medveczky et al., 1989 ).
EPIDEMIOLOGY
A serological survey in Wisconsin revealed antibod –
ies to the virus in six of 101 wild Eastern Cottontail rab –
bits ( Spieker and Yuill, 1976 ). In experimentally infected
Eastern Cottontail rabbits, herpesvirus was not recov –
ered from feces, urine, milk, ejaculates, or conjunctival
and genital secretions, but was isolated from the oral
secretions of one of 15 infected rabbits ( Spieker and
Yuill, 1977a ). Shedding of infectious virus in oral secre –
tions, unrelated to age, sex, or season, was demon –
strated in naturally infected Eastern Cottontail rabbits
(Hinze and Lee, 1980 ). The tonsils are the likely site of
persistent infection and virus is released into the oral
cavity. Transplacental transmission of virus was not
found ( Spieker and Yuill, 1977a ). Transmission by insect
vectors was not detected ( Spieker and Yuill, 1977b ).
Repeated attempts to infect domestic New Zealand
White rabbits have met with failure (Hinze, 1971a). Only
rabbits of the genus Sylvilagus appear to be susceptible.
CLINICAL SIGNS
Inoculation of Eastern Cottontail rabbits by various
routes results in a chronic low-grade viremia that per –
sists, in most instances, for the life of the rabbit ( Hinze,
1971b; Hinze and Chipman, 1972; Hinze and Wegner,
1973 ). Persistently infected rabbits have a pronounced
lymphocytosis, with differential lymphocyte counts
of up to 95% compared to 50–60% in normal rabbits
(Hinze, 1969 ).

II. RABBITS
14. VIRAL DISEASES
380
PATHOLOGY
The lymphoproliferative lesions in Eastern Cottontail
rabbits occur in the lymphoid and other organ systems
(Hesselton et al., 1988; Hinze, 1969, 1971b; Hinze and
Wegner, 1973 ). Extensive infiltration of various tissues,
commonly the kidneys, liver, lungs, and myocardium,
with immature, actively proliferating lymphocytes occurs
6–8 weeks after experimental inoculation. The experimen –
tally induced lymphoproliferative disease varies from
benign lymphoid hyperplasia to lesions consistent with
malignant lymphoma. Juvenile rabbits are affected more
frequently and severely than adult rabbits.
DIAGNOSIS
Natural infection with Leporid herpesvirus 1 in
Eastern Cottontail rabbits may result in leukocytosis,
splenomegaly, and lymphadenopathy. Virus can be
isolated from the oral cavity and circulating lympho –
cytes of infected rabbits cocultured with rabbit kidney
cells ( Hinze and Lee, 1980 ). Antibodies can be detected
by indirect immunofluorescence and plaque reduction
assays ( Spieker and Yuill, 1976; Yang et al., 1990 ).
Leporid Herpesvirus 2
HISTORY
Nesburn (1969) isolated Leporid herpesvirus 2 (syn –
onyms Herpesvirus cuniculi, virus III of rabbits, Leporid
herpesvirus 3) from primary kidney cell cultures from
Oryctolagus cuniculus and named it Herpesvirus cuniculi.
This may represent a reisolation of virus III of rabbits
(Rivers and Tillett, 1923 ), but, as the original isolate was
not available, a direct serological comparison could not
be made. Comparison of the reported characteristics of
virus III with Herpesvirus cuniculi, however, indicates
that they are identical ( Nesburn, 1969 ). Virus III was iso –
lated during attempts to find the etiologic agent of vari –
cella (chicken pox). When inoculated into rabbits, the
agent induced fever, exanthema, skin vesicles, corneal
lesions, and intranuclear inclusions reminiscent of vari –
cella infection ( Rivers and Tillett, 1923 ). Convalescent
sera from human varicella patients failed to inactivate
the virus ( Rivers and Tillett, 1924a ). A similar virus was
isolated from serially passaged normal rabbit testicular
tissue during investigations of the etiology of rheumatic
fever ( Miller et al., 1924 ). McCartney isolated the virus
while working on scarlet fever in England ( Topacio and
Hyde, 1932 ), and the agent has also been isolated by
Doerr in Switzerland ( Andrewes, 1928 ).
ETIOLOGY
Nesburn (1969) showed that the viral isolate pos –
sesses the physical, chemical, cytopathic, histologi –
cal, and electron microscopic characteristics of a
herpesvirus. The virus can be propagated in primary or established cell lines of rabbit origin as well as in
African green monkey kidney cells. The virus is now
designated Leporid herpesvirus 2 ( Roizman, 1982 ).
EPIDEMIOLOGY
Rivers and Tillett (1924a) found that four of 20 rab –
bits possessed antibodies to the virus and that 15% of
200 rabbits were immune to infection. Andrewes (1928) ,
in England, found 98% of 377 experimental rabbits sus –
ceptible to the virus. He concluded that the virus was
probably endemic in some rabbit colonies. Topacio and
Hyde (1932) , in Maryland, found that 17% of 76 rabbits
were immune to infection. They suggested that older
bucks, resistant to infection, were carriers of the virus.
Neutralizing antibodies to the virus were detected in 6%
of 196 rabbits from Connecticut and Maryland ( Swack
and Hsiung, 1972 ). Nesburn (1969) was unable to reiso –
late the virus from over 100 batches of primary rabbit
kidney cell cultures prepared since initial isolation of
the virus. Experimentally, virus was recovered from the
blood of rabbits more than 100 days after inoculation
(Swack and Hsiung, 1972 ). Appreciable titers of virus
were detected in leukocytes, spleen, liver, lungs, and sali –
vary glands, whereas the concentration of virus in kid –
neys was variable. However, transmission of the virus by
direct contact or transplacentally failed to occur.
CLINICAL SIGNS
All reported isolations of virus have been from
apparently normal rabbits, and no naturally occurring
disease has been attributed to infection with the virus.
However, a viral agent, presumably a herpesvirus, was
recovered from the nares of rabbits with respiratory dis –
ease ( Renquist and Soave, 1972 ). Experimentally, intra –
dermal inoculation of rabbits with virus resulted in
pronounced erythema at the site of inoculation after 4–7
days, which usually disappears within 2 weeks ( Rivers
and Tillett, 1924a ). Occasionally, generalized reactions
with anorexia, diarrhea, emaciation, fever, and skin ves –
icles have been reported. Intradermal inoculation may
result in erythematous papules, whereas corneal scarifi –
cation caused swelling and vesiculation of corneal cells
(Topacio and Hyde, 1932 ). Intratesticular inoculation
of rabbits results in acute orchitis and fever within 3–4
days ( Andrewes, 1928 ). Intramuscular and subcutane –
ous inoculation of rabbits failed to induce clinical signs;
however, corneal scarification with virus resulted in
mild punctate keratitis ( Nesburn, 1969 ). Similarly, intra –
peritoneal inoculation of virus did not induce clinical
disease ( Swack and Hsiung, 1972 ).
PATHOLOGY
No gross pathological changes have been observed
in internal organs of experimentally infected rabbits.

381 DNA VIRUS INFECTIONS
II. RABBITSMicroscopically, testes, skin, and cornea show edema
and an intense mononuclear leukocyte infiltration. Large
eosinophilic intranuclear inclusions, typical of herpes –
viruses, characteristically occur in corneal epithelium,
interstitial cells of the testicles, and in endothelial leuko –
cytes of the skin ( Rivers and Tillett, 1924b ). Severe myo –
carditis with typical herpesviral inclusions occurred in
rabbits inoculated intracardially with virus ( Pearce, 1950,
1960 ). The absence of reported cases could be attributed
to the relative rarity of disease or to lack of intensive etio –
logic investigation of cases which do occur.
DIAGNOSIS
Pathological findings described above including
herpesviral inclusions in domestic rabbits ( Oryctolagus
cuniculus ) are suggestive of Leporid Herpesvirus 2.
PCR, in situ hybridization, and immunohistochemi –
cal labeling would be necessary to separate Leporid
Herpesvirus 2 from the Alphaherpesvirinae described
below ( Gruber et al., 2009; Jin et al., 2008a; Weissenböck
et al., 1997 ).
CONTROL
Both herpesvirus 1 of Eastern Cottontail rabbits and
herpesvirus 2 of domestic rabbits possess the capacity to
persist in infected hosts as a subclinical infection. Such
infections, if not recognized, could result in considerable
confounding in experimental studies, especially if condi –
tions cause recrudescence of latent infections.
Leporid Herpesvirus 4
HISTORY
Leporid herpesvirus 4 (LHV 4) was isolated from a
naturally occurring outbreak with high morbidity and
mortality affecting miniature and crossbred domestic
European rabbits ( Orcytolagus cunniculi ) in a rabbitry
in Alaska ( Jin et al., 2008a ). Two other outbreaks, with
similar clinical signs and pathology, have been reported
in North America, although the virus was not identified
(Onderka et al., 1992; Swan, 1991 ).
ETIOLOGY
Electron microscopy of LHV 4 demonstrated capsid
and core particles similar in size and structure to Human
herpesvirus 1 . The genome of LHV 4 is 120–130 kbp and
partial sequences compared to other Alphaherpesvirinae
demonstrated the closest homology by nucleotide iden –
tity with Bovine herpesvirus 2 (Jin et al., 2008b ). Bovine
herpesvirus 2 was originally isolated from cattle in Africa,
but has been found in a variety of domestic and wild
ruminants ( Borchers et al., 2002; Kálmán and Egyed,
2005 ). LHV 4 infects rabbit skin and Vero (monkey kid –
ney) cells with cytopathic effects similar to Human herpes –
virus 1 (Jin et al., 2008a ).EPIDEMIOLOGY
LHV 4 was isolated from an outbreak of acute dis –
ease in an outdoor rabbitry in Alaska ( Jin et al., 2008a ).
The mode of herpesvirus introduction in this outbreak
was not determined as there had been no travel or
introduction of new rabbits. However, feral domestic
European rabbits ( Oryctolagus cuniculus ) and Snowshoe
hares ( Lepus americanus ) were in the area. In addition,
the outbreak occurred in July and August, when mos –
quito and biting fly activity is typically high. A second
outbreak occurred in the same rabbitry the following
year in the spring and summer, although it could not
be determined if this was due to a re-introduction or re-
activation of a chronic infection ( Jin et al., 2008b ). Two
outbreaks of disease caused by herpes-like viruses that
were not identified were reported in Canada ( Onderka
et al., 1992; Swan, 1991 ). The clinical signs and pathol –
ogy of those outbreaks were similar to the Alaska
outbreak and may suggest that there is an animal res –
ervoir in northwestern North America of this Alpha
herpesvirus.
CLINICAL SIGNS
Clinical signs reported in the naturally occurring out –
breaks included conjunctivitis and periocular swelling
with purulent ocular discharge, ulcerative dermatitis, pro –
gressive weakness, anorexia, respiratory distress, torticol –
lis, ataxia, weight loss, diarrhea, bruxism, genital swelling,
and abortion ( Jin et al., 2008a ). Morbidity was approxi –
mately 50%, with 29% mortality . Some rabbits died acutely
with anorexia, the only sign observed prior to death.
Recovered virus administered intranasally or by the cor –
nea to New Zealand White rabbits caused severe acute
disease within 4 days of inoculation with one out of four
affected animals being euthanized due to severity of the
disease. The remaining rabbits recovered. Acute mortality
associated with unidentified herpesviruses was reported
in four adult Oryctolagus cuniculus in two rabbitries in
Canada ( Onderka et al., 1992; Swan, 1991 ). Acute death
without clinical signs occurred in some rabbits as well as
conjunctivitis, nasal discharge, edematous eyes and faces.
Herpesviruses were recovered on rabbit kidney cells and
experimental inoculation of rabbits with the virus repro –
duced the disease syndrome ( Onderka et al., 1992; Swan,
1991 ). However, it is unclear if the herpesviruses respon –
sible for these outbreaks are Gamma herpesviruses or
Alpha herpesviruses related to LHV 4.
PATHOLOGY
Lesions included severe dermal and subcutaneous
hemorrhage, edema, and necrosis with vascular necro –
sis and thrombosis ( Jin et al., 2008a, 2008b ). There was
a mixed infiltrate of macrophages, lymphocytes, plasma
cells with fewer heterophils. Necrosis and hemorrhage

II. RABBITS
14. VIRAL DISEASES
382
were found within the ventricular walls, spleen, lymph
nodes, and lungs. Areas of fibrin deposition were pres –
ent in the spleen and lung. Intranuclear inclusions were
found within the affected skin, subcutis, and ventricu –
lar walls, but not in the spleen, lungs, or lymph nodes.
Similar lesions were seen following experimental infec –
tion ( Jin et al., 2008b ).
DIAGNOSIS
Typical lesions in the skin, spleen, lymph nodes,
lungs, and heart with demonstrative intranuclear inclu –
sions are suggestive of LHV 4 infection. DNA extracted
from most tissues, including appendix, sciatic nerve,
kidney, adrenal gland, spleen, liver, lung, lymph node,
salivary gland, tonsil, and brain from an inoculated rab –
bit that died in the acute phase of the disease was spe –
cific for LHV 4 ribonuclease reductase gene by PCR ( Jin
et al., 2008b ). At 14 days after inoculation, the eye, tri –
geminal ganglion, and tonsil were positive for LHV 4
by PCR analysis. In the second outbreak at the rabbitry
in Alaska, the eye of an affected rabbit was positive for
LHV 4 by PCR.
CONTROL
As latent infection is typical of Alpha herpesviruses,
control measures are difficult. The rabbitry involved in
the Alaska outbreak was depopulated after the second
outbreak (Jin et al., 2008).
Human Herpesvirus 1 (HHV-1)
Herpesviruses of other species, including HHV-1, can
cause encephalitis in rabbits experimentally inoculated
(Chowdhury et al., 2000; Seto et al., 1995 ) and horizon –
tal transmission of HHV-1 between rabbits has been
reported ( Kaplan, 1969 ). Spontaneous cases of HHV-1
encephalitis presumably transmitted from humans to
pet domestic European rabbits have also been reported
(Grest et al., 2002; Gruber et al., 2009; Müller et al., 2009;
Weissenböck, et al., 1997 ). In each of these reports, rab –
bits with HHV-1 encephalitis were in close contact with
a person with herpes labialis before they developed ill –
ness. The rabbits presented with neurological signs
including anorexia, tonic-clonic spasms, circling, incoor –
dination, ataxia, seizures, and opisthotonus. One rabbit
also had a history of epiphora of the left eye, conjuncti –
vitis, bruxism, and hypersalivation ( Müller et al., 2009 ).
Pathological findings included non-suppurative menin –
goencephalitis and large, eosinophilic intranuclear inclu –
sion bodies in the neurons and glial cells. Diagnosis was
confirmed by immunohistochemical labeling with poly –
clonal antibodies specific for HHV-1 ( Grest et al., 2002;
Gruber et al., 2009; Weissenböck et al., 1997 ), in situ
hybridization using a DNA probe for HHV-1 ( Gruber
et al., 2009; Müller et al., 2009; Weissenböck et al., 1997 ), or PCR specific for the UL42 gene of HHV-1 ( Grest
et al., 2002; Gruber et al., 2009; Weissenböck et al., 1997 ).
Control measures would include preventing close con –
tact between humans with active herpetic lesions and
rabbits.
Papilloma and Polyoma Viruses
The family Papillomaviridae contains viruses in the
genus Kappapapillomavirus which cause papillomas
(warts) in various animals and includes two species of
rabbit viruses, the Cottontail rabbit papillomavirus and
the Rabbit oral papillomavirus (Fauquet et al., 2005 ). The
family Polyomaviridae contains one genus, Polyomavirus ,
which includes the species Rabbit kidney vacuolating virus .
Cottontail Rabbit Papillomavirus
HISTORY
The Cottontail rabbit or Shope papillomavirus was
recognized as a transmissible agent by Shope and Hurst
(1933) while they were attempting to define the etiol –
ogy of wart-like tumors on Eastern Cottontail rabbits
(Sylvilagus floridanus ) in the Midwestern United States.
They demonstrated the filterability of the infectious
agent, transmitted it to other Eastern Cottontail rabbits
as well as domestic European rabbits, and described
the histopathology of the disease ( Shope, 1935 ). Rabbit
papillomatosis was initially considered a natural benign
disease of Sylvilagus rabbits only; however, spontane –
ous outbreaks of papillomatosis in Oryctolagus cuniculus
indicate that the disease has greater relevance ( Hagen,
1966 ). Soon after discovery of the virus, it was also
shown to cause malignant neoplasms histologically
resembling squamous cell carcinomas ( Kidd and Rous,
1940; Rous and Beard, 1934, 1935 ). This was the first
recognition of an oncogenic virus in mammals. The dis –
ease has served as a model system to elucidate the role
of viruses in the pathogenesis of cancer in humans and
animals ( Evans and Thomsen, 1969; Evans et al., 1962a,
1962b; Georges et al., 1984; Kreider and Bartlett, 1981,
1985; Smith and Campo, 1985 ) and potential therapeutic
interventions ( Gambhira et al., 2007 ).
ETIOLOGY
The virus is the type species of the genus
Kappapapillomavirus (formerly Papillomavirus ) of the fam –
ily Papillomaviridae (Fauquet et al., 2005 ) and possesses
the characteristic circular DNA genome, icosahedral
structure, and other chemical and physical properties of
this family ( Gross, 1983; Kass and Knight, 1965; Murphy
et al., 1981 ). The complete genome contains 7863 nucleo –
tides ( Giri et al., 1985 ) and consists of ten genes. All of
the papilloma viruses have two capsid proteins (major
L1 and minor L2) and although the C terminus of L1 is

383 DNA VIRUS INFECTIONS
II. RABBITSexposed on the surface of the viron, suggesting it may
have an important role in immune responses, both pro –
teins encode virus-neutralizing epitopes ( Campo, 2003 ).
The virus is antigenically distinct from other members of
the genus Kappapapillomavirus. Nuclear ( Yoshida and Ito,
1968 ) and cytoplasmic ( Ishimoto et al., 1970 ) viral anti –
gens have been demonstrated by immunofluorescence
in virus-inoculated normal skin cells and virus-induced
papillomas. However, the significance of these findings
is difficult to interpret since controls for rabbit kidney
vacuolating virus, a passenger virus ( Hartley and Rowe,
1964 ), were usually not included ( Kreider and Bartlett,
1981 ). The virus can be maintained by serial propagation
in the skin of Sylvilagus rabbits, inoculated intracutane –
ously or scarified with virus ( Shope and Hurst, 1933 ).
Intramuscular inoculation does not result in clinical papil –
lomatosis. Dogs, cats, pigs, goats, rats, mice, and guinea
pigs are refractory to the virus. The skin of embryonic rats
is susceptible to the virus, and typical papillomas develop
following inoculation ( Greene, 1953b ); however, few
papillomas develop, and they regress unless the host is
immunosuppressed ( Kreider et al., 1971 ). Cellular prolif –
eration or neoplastic transformation by the virus has been
reported in rabbit skin cell cultures ( Coman, 1946 ), skin
cultures derived from neonatal domestic European rab –
bits ( DeMaeyer, 1962 ), and explants of embryonic rabbit
skin ( Greene, 1953a ). Transplantation of transformed skin
cultures into rabbits resulted in the formation of papillo –
mas ( Coman, 1946; Kreider, 1963 ). (see also Chapter 16).
EPIDEMIOLOGY
Papillomatosis occurs most frequently as a natural
disease of Sylvilagus rabbits in the Midwestern United
States extending from Minnesota and North Dakota to
Texas ( Gross, 1983; Hagen, 1966 ). The virus has appar –
ently not become established in the eastern states. The
only natural outbreaks of disease in domestic European
rabbits were in southern California, suggesting that the
virus is present in wild Sylvilagus rabbits of the area,
and spreads to domestic European rabbits by arthropod
vectors ( Hagen, 1966 ). The disease in Sylvilagus rabbits
also occurs on coastal islands in Washington, in a popu –
lation of rabbits introduced from Kansas ( Lancaster and
Olson, 1982 ).
The Sylvilagus rabbit is the natural host, but the
domestic European rabbit is also susceptible to the virus
(Shope, 1935 ). Natural outbreaks have been reported in
commercial rabbitries ( Hagen, 1966 ). The Black-tailed
jackrabbit ( Lepus californicus ) is also susceptible to the
virus ( Beard and Rous, 1935 ). Infection of the skin of
domestic European rabbits results in the formation of
papillomas essentially devoid of infectious virus ( Shope,
1935 ). Papillomas, primary carcinomas, and metastases
from domestic European rabbits have about 10–100 cop –
ies of viral DNA, whereas papillomas from Sylvilagus rabbits have 2400–8800 copies ( Stevens and Wettstein,
1979 ). Papillomas from domestic European rabbits can be
serially transferred, but only small amounts of infectious
virus are typically demonstrated ( Gross, 1983; Hu et al.,
2007; Selbie and Robinson, 1947; Shope, 1935 ), indicating
that domestic European rabbits are not a source of virus
for arthropod vectors.
Infection of Sylvilagus rabbits, Black-tailed jackrabbits,
and Snowshoe hares ( Lepus americanus ) induces lesions
containing a high concentration of virus ( Lancaster and
Olson, 1982; Syverton et al., 1950 ). Contact transmission
of papillomatosis may occur, but transmission of the
virus by the rabbit tick ( Haemaphysalis leporis-palustris ) is
probably the most common natural mode ( Larson et al.,
1936 ). Transmission by mosquitoes and reduviid bugs
has been demonstrated experimentally ( Dalmat, 1958 ). In
commercial rabbitries, the mosquito may be the principal
vector between wild Sylvilagus and domestic European
rabbits ( Hagen, 1966 ). This hypothesis is strengthened by
the observation that in natural cases of disease in domes –
tic European rabbits, lesions were confined to the rela –
tively hairless parts of the body around the eyes, ears,
and anus, areas where mosquitoes are more apt to feed.
Nematodes may be involved in the natural transmission
of virus ( Rendtorff and Wilcox, 1957 ). Experimentally,
papillomas were induced when papilloma virus and lar –
vae of the nematode Nippostrongylus muris were applied
to rabbit skin, but not by virus or nematode larvae alone.
See Chapter 16 for further information.
CLINICAL SIGNS
Papillomatosis of wild Sylvilagus rabbits is charac –
terized by the presence of horny warts, usually on the
neck, shoulders, or abdomen. The warts begin as red
raised areas at the site of infection, grow to become typi –
cal papillomas with rough rounded surfaces, and may
later develop into large, keratinized horny growths
(Shope and Hurst, 1933 ). The virus was initially believed
to cause only transient papillomatosis, but it was later
shown that in naturally infected Sylvilagus rabbits papil –
lomas may become malignant squamous cell carcinomas
(Syverton and Berry, 1935 ). This phenomenon was later
shown to be a relatively frequent occurrence in both nat –
urally and experimentally infected Sylvilagus rabbits, as
25% of infected rabbits developed carcinomatous lesions
following infection ( Syverton et al., 1950 ). In approxi –
mately 35% of naturally infected rabbits, papillomas
regress within 6 months after infection. In natural out –
breaks of papillomatosis of domestic European rabbits in
southern California ( Hagen, 1966 ), papillomas most com –
monly occurred on the eyelids and ears. Experimentally
induced papillomas develop more slowly in domestic
European rabbits than in Sylvilagus rabbits, reach a sta –
tionary phase, and then occasionally regress (Syverton,
1952).

II. RABBITS
14. VIRAL DISEASES
384
Regression occurs depending on the interaction of
the genotype of the virus and host immunity ( Hu et al.,
2002, 2005 ). A variant virus that contains the regressive
E6 gene has a high rate of regression ( Hu et al., 2002 ).
Certain MHC haplotypes have been associated with
regression or persistence in domestic European rabbits
(Salmon et al., 2000 ). In one study, papillomas persisted
when a regressive viral strain was used if rabbits were
immunosuppressed with cyclosporine A treatment, but
regression also varied with the strain of rabbit as more
papillomas regressed in outbred New Zealand White
rabbits than inbred EIII/JC rabbits ( Hu et al., 2005 ).
Once immunosuppression was removed, most papil –
lomas regressed in outbred rabbits (67–89%), whereas
only 13% regressed in inbred rabbits, highlighting the
importance of multiple factors in persistence of pap –
illomas. Rous and Beard (1934, 1935) recognized the
malignant potential of rabbit papillomavirus when
they demonstrated that intramuscular inoculation
of papillomatous tissue into domestic European rab –
bits resulted in invasive squamous cell carcinomas. In
experimentally infected domestic European rabbits,
75% developed carcinomas if kept longer than 6 months
(Syverton, 1952 ). Sylvilagus rabbits have a three-fold
lower incidence of carcinomas than European rabbits
(Rous and Beard, 1935; Syverton et al., 1950 ).
PATHOLOGY
The warts which develop following infection in
Sylvilagus and domestic European rabbits are typical
papillomas. The malignant tumors which arise from
papillomas are squamous cell carcinomas. Metastasis to
regional lymph nodes, particularly the axillary nodes,
is common (Kreider and Bartlet, 1981) and about 25%
of rabbits that succumb have pulmonary metastases. In
addition, amyloid deposition in renal glomeruli, hepatic
sinusoids and splenic red pulp is present in the majority
of rabbits.
DIAGNOSIS
Cottontail rabbit papillomatosis is diagnosed clinically
by the characteristic skin tumors, which never occur in
the mouth, and may be confirmed by histopathological
examination. A survey of tumors in pet rabbits submit –
ted to the surgical biopsy service of a veterinary school in
Pennsylvania over a 16-year period revealed that Shope
papilloma was diagnosed in two tumors, both on the
ears, from 179 rabbits ( von Bomhard et al., 2007 ). No cell
culture system is available for routine isolation of virus
(Lancaster and Olson, 1982 ).
CONTROL
The endemic disease of Sylvilagus rabbits is of little
economic significance, and thus no prophylactic meth –
ods have been developed. Because natural infection
occurs in domestic European rabbits ( Hagen, 1966 ), the adoption of control methods may become necessary. In
areas where the disease is endemic in wild Sylvilagus
rabbits, where arthropod vectors are present, and where
outdoor rabbit husbandry is practiced, arthropod con –
trol would appear to be a logical approach. Rabbits
can be immunized by two intraperitoneal inoculations
with glycerinated rabbit papilloma suspensions ( Shope,
1937 ). Domestic rabbits with experimentally induced
papillomas resist challenge with virus derived from
papillomas of cottontail rabbits ( Hagen, 1966 ). A tumor-
specific vaccine composed of allogenic tumor cells
increased the regression rate of papillomas ( Evans et al.,
1962a ).
Rabbit Oral Papillomavirus
HISTORY
Oral papillomatosis of rabbits was recognized as
a distinct viral disease of domestic European rabbits
(Oryctolagus cuniculus ) by Parsons and Kidd (1936) .
They found a 17% prevalence of small papillomas in the
mouths of rabbits of several breeds in New York City.
The lesions were usually confined to the ventral sur –
face of the tongue. They transmitted the virus to both
domestic European and Sylvilagus rabbits and by cross-
immunity tests and tissue-susceptibility studies dem –
onstrated that it was distinct from the Cottontail rabbit
(Shope) papillomavirus. They also showed that virus
is frequently present in the mouths of rabbits without
lesions and that tattooing or licking of tar stimulated
lesion development in such carrier rabbits ( Parsons and
Kidd, 1943 ). A spontaneous outbreak in New York of
rabbit oral papillomatosis, involving several breeds of
domestic European rabbits, was described ( Weisbroth
and Scher, 1970 ). New Zealand White rabbits with oral
papillomas were reported in Illinois ( Sundberg et al.,
1985 ) and Mexico ( Dominguez et al., 1981 ).
ETIOLOGY
The virus was partly characterized by Parsons
and Kidd (1943) and was later included in the genus
Kappapillomavirus of the family Papillomaviridae
(Fauquet et al., 2005 ). The virus consists of 7565 nucle –
otides and has the greatest amino acid sequence iden –
tity to Cottontail rabbit papillomavirus , although the
greatest area of homology was only 68% ( Christensen
et al., 2000 ). However, the virus is immunologically
distinct from the Cottontail rabbit papillomavirus and
naturally infects only leporids ( Parsons and Kidd,
1936, 1943 ). Nuclear viral replication, characteristic
of the Papovaviridae , has been demonstrated ( Rdzok
et al., 1966; Richter et al., 1964 ). Neonatal hamsters,
inoculated subcutaneously with a tumor suspension
from naturally infected rabbits, developed fibromas
(Sundberg et al., 1985 ). The virus has not been propa –
gated outside of susceptible host animals.

385 DNA VIRUS INFECTIONS
II. RABBITSEPIDEMIOLOGY
The virus probably spreads by direct contact, and
lesion development may require oral trauma for viral
entry. Coarse feed, maloccluded teeth, or other oral
irritants may serve as the inciting event ( Parsons and
Kidd, 1943; Weisbroth and Scher, 1970 ). Experimentally
induced lesions appear 9–38 days after inoculation
(Parsons and Kidd, 1936 ), but the incubation period of
the natural disease is unknown. The disease generally
occurs in rabbits 2–18 months old ( Sundberg et al., 1985;
Weisbroth and Scher, 1970 ).
CLINICAL SIGNS
Oral papillomatosis is a benign disease character –
ized clinically by small discrete whitish growths on
the ventral surface of the tongue. The early lesions are
sessile, later become rugose or pedunculated, and ulti –
mately ulcerate ( Parsons and Kidd, 1936; Sundberg
et al., 1985; Weisbroth and Scher, 1970 ). The lesions are
seldom more than 5 mm in size and 4 mm in thickness
and usually substantially smaller. Papillomas have been
known to persist for as long as 145 days, but usually
disappear in weeks ( Parsons and Kidd, 1936 ). Lesions
rarely occur elsewhere in the mouth and never on the
body. However, in a Flemish Giant pet rabbit in New
Zealand, virus recovered from lesions on the nictitating
membrane and lower eyelid had 99.3% homology with
Rabbit oral papillomavirus (Munday et al., 2007 ).
PATHOLOGY
The lesions are microscopically typical papillomas
(Parsons and Kidd, 1943; Rdzok et al., 1966; Richter et al.,
1964; Sundberg et al., 1985; Weisbroth and Scher, 1970 ).
Cells in the stratum spinosum contain basophilic intra –
nuclear inclusions ( Dominguez et al., 1981; Sundberg
et al., 1985 ).
DIAGNOSIS
The disease is diagnosed by typical lesions occurring
only in the mouth, in contrast to rabbit papillomatosis, in
which lesions are observed only on the skin. The lesions
are typical papillomas, and papillomavirus antigens can
be detected in cells of the stratum spinosum by the per –
oxidase antiperoxidase technique ( Sundberg et al., 1985 ).
CONTROL
Rabbits recovering from disease are resistant to rein –
fection but are susceptible to Cottontail rabbit (Shope)
papillomavirus .
Rabbit Kidney Vacuolating Virus
The Rabbit kidney vacuolating virus was isolated in
primary rabbit kidney cell cultures from papillomas of
Eastern Cottontail rabbits ( Sylvilagus floridanus ) collected
in Kansas ( Hartley and Rowe, 1964 ). The virus causes vacuolar cytopathic effects in cell cultures. The virus
resembles the Cottontail rabbit papillomavirus but is a dis –
tinct virus. It does not produce papillomas when inocu –
lated into rabbits and does not immunize rabbits against
Cottontail rabbit papillomavirus . The virus is slightly smaller
than the viruses of the genus Papillomavirus and resembles
Polyomavirus in size, morphology , and DNA composition
(Crawford and Follett, 1967 ). The virus has been classified
as a member of the genus Polyomavirus within the fam –
ily Polyomaviridae (Fauquet et al., 2005 ). Ultrastructurally ,
the virus is a typical Polyomavirus in its replication pat –
tern ( Chambers et al., 1966 ). The virus is not pathogenic
for either domestic European or Eastern Cottontail rab –
bits or any other animal species. Inoculation of both neo –
natal and adult domestic and Eastern Cottontail rabbits
by several routes did not induce disease ( Hartley and
Rowe, 1964 ). Antibodies to the virus have been found in
wild Sylvilagus rabbits in Kansas and Maryland, but not in
domestic European rabbits ( Hartley and Rowe, 1964 ). The
virus thus appears to be a fairly common non-pathogenic
virus of Sylvilagus rabbits, causing only latent infections.
Intranuclear inclusions consistent with rabbit kidney
vacuolating virus have been found as an incidental find –
ing in New Zealand White rabbits ( Percy and Barthold,
2007 ). Although a contaminant of rabbit papillomas, the
virus appears to have no role in the pathogenesis of pap –
illomatosis ( Goldman et al., 1972; Ito et al., 1968; Kreider
and Bartlett, 1981 ).
Adenovirus Infections
Adenovirus infections of rabbits, spontaneous and
experimental, have been reported. While the taxonomy
of the spontaneous adenoviruses found in rabbits has not
been determined, the human, swine, and bovine adeno –
viruses described belong to the genus Mastadenovirus . An
adenovirus was isolated in Hungary from the spleen, kid –
neys, lungs, and intestines of rabbits, 6–8 weeks old, with
diarrhea ( Bodon et al., 1979 ). The virus was detected in pri –
mary rabbit kidney cultures stained with acridine orange
although no cytopathic effect was observed. The virus
failed to replicate in pig kidney , calf kidney , calf testicle, or
human embryonic lung cells. Antisera to several swine and
bovine adenoviruses failed to neutralize the virus, whereas
a partial antigenic relationship to human adenoviruses was
demonstrated by complement fixation and immunodif –
fusion tests. The virus agglutinated rabbit but not human
erythrocytes. A serological survey of 30 Oryctolagus cunic –
ulus from four rabbitries in Quebec revealed that three
rabbits from three colonies had antibodies to bovine ade –
novirus type 1 ( Descoteaux et al., 1980 ).
Experimental inoculation of rabbits with human adeno –
virus type 5 resulted in no clinical response but induced a
persistent viral infection in lymphoid tissues for as long as
1 year ( Reddick and Lefkowitz, 1969 ). Human adenovirus
type 5 infects rabbit corneas following either intrastromal

II. RABBITS
14. VIRAL DISEASES
386
injection or topical application to damaged cornea result –
ing in acute blepharoconjunctivitis, iritis, and corneal
edema which peaks at 13 days after infection ( Gordon et
al., 1992; Trousdale et al., 1995 ). Other serotypes of human
adenovirus induce disease as well, although adenovi –
rus type 5 is commonly used as an animal model to test
therapeutic approaches to adenovirus-induced kerato –
conjunctivitis ( Romanowski et al., 1998 ). Recombinant
adenoviruses have been used successfully to infect rab –
bits in many gene transfer protocols ( Kozarsky et al, 1996;
Schneider et al., 1999 ).
Parvovirus Infections
A virus that induced cytopathic effects in primary
rabbit kidney cell cultures was isolated from the feces
of a rabbit ( Oryctolagus cuniculus ) inoculated with
Herpesvirus cuniculi in a laboratory in Japan ( Matsunaga
et al., 1977 ). The virus had the morphological, physi –
cal, and chemical properties of a parvovirus. The
various proteins of the virus have been characterized
(Matsunaga and Matsuno, 1983 ). Subsequently, parvo –
virus was recovered from kidney cells of neonatal rab –
bits in the United States ( Metcalf et al., 1989 ). These
viruses are of the species Lapine parvovirus , genus
Parvovirus , family Parvoviridae (Fauquet et al., 2005 ).
Serological surveys have identified positive rabbitries
in Japan, the United States, and Europe. Among 90 rabbits
from a commercial source in Japan, 42 (47%) had hemag –
glutination-inhibiting antibody to the virus ( Matsunaga et
al., 1977 ). Of 46 rabbit sera, collected from various sources
in the United States, 75% had antibodies detected by
immunofluorescence or hemagglutination inhibition assays
(Metcalf et al., 1989 ). In Switzerland, over 70% of 132 rabbit
sera from various commercial breeding colonies had anti –
bodies in the hemagglutination inhibition test ( Metcalf et
al., 1989 ). Experimental inoculation, orally or intravenously ,
induced anorexia, listlessness, and catarrhal enteritis in
1-month-old rabbits ( Oryctolagus cuniculus ) (Matsunaga
and Chino, 1981 ). Pathologic changes included catarrhal
enteritis with hyperemia, exfoliation of the small intestinal
epithelial cells, and increased secretion of intestinal mucus.
Virus was detected in feces, small intestine, liver, pancreas,
spleen, appendix, and mesenteric lymph node. Rabbits
developed hemagglutination-inhibiting antibodies.
RNA VIRUS INFECTIONS
Rotavirus Infections
Rotavirus
HISTORY
Rotavirus was initially isolated from rabbits with
diarrhea by Bryden et al. (1976) in England. Virus was recovered from both sporadic cases and outbreaks of
diarrhea in weanling rabbits as well as from healthy rab –
bits. Subsequently, rotavirus was isolated from young
rabbits with diarrhea in Japan ( Sato et al., 1982 ), Europe
(Castrucci et al., 1985; Eaton, 1984; Peeters et al., 1982,
1984 ), and the United States ( DiGiacomo and Thouless,
1986; Schoeb et al., 1986 ). Various serological surveys
have since revealed that rotavirus infection is wide –
spread in domesticated European rabbits ( Oryctolagus
cuniculus ).
ETIOLOGY
Rabbit rotaviruses belong to the family Reoviridae ,
genus Rotavirus (Estes and Kapikian, 2007; Fauquet
et al., 2005 ). They are 75-nm icosahedral viruses with a
triple-layered protein capsid that contains 11 segments
of double-stranded RNA, each of which codes for a
protein. The sixth gene (VP6) codes for an inner capsid
protein which contains the subgroup specificity. The
fourth (VP4) and seventh genes (VP7), code for outer
capsid proteins that are the viral neutralization anti –
gens. VP4 is the viral hemagglutin and must be cleaved
by a protease, e.g., trypsin, to yield VP5 and VP8 before
the virus can infect cells ( Matsui et al., 1989; Ramig and
Ward, 1991 ).
The rotaviruses are classified serologically into seven
groups (A to G). Groups A to C are found in animals
and humans, whereas groups D to G are found only
in animals. The rabbit rotaviruses described belong
to group A ( Tanaka et al., 1988; Thouless et al., 1986 ).
Within each serogroup, rotaviruses are classified into
serotypes, defined by reactivity against the outer capsid
proteins VP4 (P type) and VP7 (G type). Since the gly –
coprotein VP7 appears to be immunodominant, classi –
fication of VP7 serotypes is more advanced and within
group A, 15 serotypes (1–15) have been identified. All
rabbit rotaviruses recovered to date have been sero –
type 3 (G3) ( Tanaka et al., 1988; Thouless et al., 1986 ).
Serotype 3 contains the largest and most diverse num –
ber of rotaviruses and is common to humans and sev –
eral other animals. This reflects the genetic reassortment
that occurs when different viruses of the same group
simultaneously infect susceptible hosts.
Some rotavirus isolates from rabbits induce a cyto –
pathic effect in MA104 rhesus monkey kidney cells
in about 3 days ( Castrucci et al., 1985; Sato et al., 1982;
Thouless et al., 1986 ), whereas other isolates demon –
strate cytopathic effect only after additional passages.
Physicochemical studies reveal that rotaviruses are
stable from pH 3.0 to 9.0 (Fauquet et al ., 2000). Virus
remains infectious for months at room temperature
on porous and non-porous environmental surfaces, in
fecal material, or if stabilized with CaCl 2. Rotaviruses
can be inactivated when heated at 50°C for 30 min and
by treatment with chlorine, formalin, phenols, and

387 RNA VIRUS INFECTIONS
II. RABBITSbetapropriolactone. Ethanol, 95%, is the most effective
disinfectant as it renders the rotavirus non-infectious by
removing the outer capsid layer.
EPIDEMIOLOGY
Serological studies indicate that rotavirus infec –
tion is widespread in domesticated European rabbits
(Oryctolagus cuniculus ). Of 91 adult rabbits from two
commercial rabbitries in Ontario, Canada, 98% had
antibodies to rotavirus (Petrie et al., 1978). Sera col –
lected from fryer rabbits at two abattoirs in Ontario,
Canada, revealed that 60% of 200 rabbits had antibod –
ies ( Percy et al., 1993 ). A survey in Tokyo prefecture,
Japan, of 39 adult rabbits revealed that 82% had anti –
bodies ( Takahashi et al., 1979 ). A more extensive survey
of ten breeding and 13 laboratory colonies in metropoli –
tan Tokyo and Ibaragi, Saitama, Kanagawa, Shizuoka,
and Nagano prefectures in Japan revealed antibodies
to rotavirus in 83% of the 23 colonies and in 81% of 160
sera ( Iwai et al., 1986 ). In Hungary, 74% of 112 sera from
five large-scale rabbit farms had rotavirus antibodies
(Kudron et al., 1982 ). In the United States, 95% of 149
sera from rabbits more than 2 months old in a commer –
cial colony in Washington had antibodies ( DiGiacomo
and Thouless, 1984 ). Rotavirus was detected in fecal
samples from rabbits in three other colonies in the
state ( DiGiacomo and Thouless, 1986 ). Antibodies
against rotavirus have also been detected in 29% of 17
sera from Eastern Cottontail rabbits (Sylvilagus florida –
nus) in Ontario, 52% of 27 sera from Snowshoe hares
(Lepus americanus ) in the Yukon, and 6% of 48 sera from
Snowshoe hares in Nova Scotia (Petrie et al ., 1978).
In colonies with endemic rotavirus infection, nearly
all adult rabbits have serological evidence of rotavirus
infection ( DiGiacomo and Thouless, 1984, 1986 ). Litters
in such colonies have transplacentally acquired mater –
nal antibodies to rotavirus at birth. In the absence of
rotavirus infection, the antibodies fall to undetectable
levels by 60 days of age. In colonies with endemic infec –
tion, shedding of rotavirus in the feces is detected in
rabbits 4–7 weeks old ( Peeters et al., 1984 ), followed by
the appearance of naturally acquired antibodies in rab –
bits greater than 6 weeks old ( DiGiacomo and Thouless,
1986 ). Subsequently, infected rabbits demonstrate anti –
bodies for long periods. Hence, in infected colonies,
rabbits usually acquire infection when maternal anti –
bodies have declined to low concentrations, which
usually coincides with weaning. Both sexes appear
equally susceptible. Rotavirus infection in domesticated
European rabbits has been reported in New Zealand
White, Dutch, and Californian rabbits, suggesting no
difference in breed susceptibility ( Bryden et al., 1976 ).
Rotavirus is shed in the feces of infected rabbits. In
a survey of 187 rabbits of mixed ages, rotavirus was
detected in 4% of fecal samples ( Petric et al., 1978 ). In another study, 11% of 18 fecal samples contained virus
(Kudron et al., 1982 ). Rotavirus was detected in 9% of
106 fecal samples from healthy rabbits, 1–2 months
old, from four rabbitries ( DiGiacomo and Thouless,
1986 ). Hence, transmission probably occurs by fecal–
oral spread. Rabbits inoculated orally with rotavirus
shed virus in the feces for 6–8 days, beginning 2–5 days
after inoculation ( Blutt et al., 2003; Castrucci et al., 1984;
Conner et al., 1988; Hambraeus et al., 1989 ; Petrie et al.,
1978; Thouless et al., 1988 ). Furthermore, uninoculated
control rabbits maintained in the same room with inoc –
ulated rabbits also acquired rotavirus infection ( Conner
et al., 1988; Thouless et al., 1988 ). As fomite transmis –
sion appeared unlikely because of the handling of unin –
oculated rabbits first, it was concluded that rabbits were
infected by airborne transmission of virus. In gastrically
inoculated rabbits, rotavirus RNA was also detected
in sera 4 days after inoculation ( Blutt et al., 2003 ).
Experimentally, rabbits were also infected with simian
rhesus rotavirus ( Ciarlet et al., 2000b ).
Recent reports suggest that rabbits may be a source
of rotaviruses for humans. Comparison of the sequence
analysis of several rotavirus genes of rabbit and human
origin revealed a close relationship with clustering of
strains in phylogenetic analyses ( De Leener et al., 2004;
Matthijnssens et al., 2006, 2009 ). However, the molecu –
lar similarity of strains should not infer interspecies
transmission without supporting epidemiologic evi –
dence of spread.
CLINICAL SIGNS
Rotavirus was initially detected and recovered from
rabbits with diarrhea by Bryden et al. (1976) . That report
included both sporadic cases and outbreaks in 5-week-
old rabbits. A spectrum in the severity of disease associ –
ated with rotavirus infection has been reported, which
is probably influenced by a synergy among various
microorganisms responsible for diarrheal diseases in
rabbits. In outbreaks of diarrhea associated with rotavi –
rus infection, rabbits 30–80 days old are usually affected
(Castrucci et al., 1985; Peeters et al., 1984; Sato et al.,
1982 ). Rabbits exhibit severe mucoid or watery diarrhea,
anorexia, and dehydration, with mortality of 60–80%. In
one outbreak, rabbits 8–12 days old had a greenish-yel –
low watery diarrhea ( Peeters et al., 1982 ). About 20% of
litters were affected, with 98% mortality within 1–2 days
after onset of signs. A similar disease was reported in a
specific pathogen-free colony in which litters 7–21 days
old exhibited watery diarrhea and lethargy ( Schoeb et al.,
1986 ). Approximately 40% of affected litters died within
2 days after onset of signs. That preweanling rabbits
were affected in both outbreaks suggests that rotavirus
had been recently introduced into the colonies.
In a comprehensive study of various infectious
agents (parasites, bacteria, and viruses) associated with

II. RABBITS
14. VIRAL DISEASES
388
diarrhea in 21 rabbitries, Peeters et al. (1984) detected
rotavirus in 35% of 130 affected rabbits. However, the
clinical signs associated with rotavirus infection con –
sisted of watery diarrhea for 2–3 days, with low mor –
tality. Hence, in endemically infected colonies, other
factors, including other infectious agents, may enhance
the pathogenicity of rotavirus ( Peeters et al., 1984 ). That
rotavirus may be only mildly pathogenic is supported
by experimental studies. In general, orogastric inocula –
tion of rabbits, 1–22 weeks old, with rotavirus did not
result in diarrhea, although some rabbits developed
soft or fluid feces for 2–4 days ( Conner et al., 1988;
Hambraeus et al., 1989; Petric et al., 1978; Thouless
et al., 1988 ). Whereas inoculation of one rotavirus strain
induced diarrhea, depression, anorexia, and mortal –
ity in rabbits ( Castrucci et al., 1984 ), this could not be
repeated ( Hambraeus et al., 1989 ). Inoculation of rabbits
of various ages revealed that partially formed to liquid
stools developed in 1-week-old rabbits but not in rab –
bits 2 weeks old ( Ciarlet et al., 1998 ). Thus, in non-
endemically infected rabbits, rotaviral disease may be
age-restricted, occurring only in neonatal rabbits.
PATHOLOGY
In weanling rabbits infected with rotavirus, the intes –
tines are markedly congested and distended, and pete –
chiae are found in the colon ( Sato et al., 1982 ). In addition
to congestion, there are mucosal hemorrhages in the
small intestine and distention of the cecum with fluid
(Castrucci et al., 1985 ). However, in both of the reports,
the presence of other infectious agents was not exam –
ined. Peeters et al. (1984) reported that, in pure rotavirus
infection, gross lesions are limited to fluid cecal contents
and swollen mesenteric lymph nodes. Histologically, the
small intestine shows moderate to severe villous atrophy,
more marked in the ileum. Apical enterocytes on the tips
of villi are swollen, rounded, and desquamating; occa –
sionally tips are denuded. The lamina propria is usually
infiltrated with lymphocytes and occasional neutrophils.
Lesions in the cecum are limited to focal areas of entero –
cyte desquamation. In outbreaks involving preweanling
rabbits, gross lesions are most pronounced in the ileum
(Peeters et al., 1982 ). Microscopically, there is villous atro –
phy and attenuation or desquamation of epithelial cells
at the apical tips of jejunal or ileal villi ( Peeters et al.,
1982; Schoeb et al., 1986 ). In some areas, the submucosa
is edematous ( Schoeb et al., 1986 ).
Experimentally, rabbits inoculated orally with rotavi –
rus have markedly congested and distended intestines
with accumulation of fluid and gas in the small intes –
tine, cecum, and colon from 2–9 days after inoculation
(Castrucci et al., 1984; Ciarlet et al., 1998; Petric et al.,
1978 ; Thouless et at., 1988). Microscopically, villi are
shortened with villus blunting, fusion, and vacuolation,
low to moderate numbers of lymphocytes and plasma cells infiltrate the villi and lamina propria of the small
intestine. There is mild lymphoid reactivity in mes –
enteric lymph nodes. Lesions are not observed in the
cecum or colon. The pathologic changes are considered
to be mediated by enterotoxin ( Ciarlet et al., 2000a ).
DIAGNOSIS
Diarrhea caused by rotavirus is diagnosed from clini –
cal signs, histopathology, detection of the virus, and
demonstration of antibodies. Clinical signs and patho –
logical findings alone are not diagnostic, although vil –
lous atrophy and degeneration and desquamation of
enterocytes at the tips of villi in the small intestine are
characteristic features. Rotaviral diarrhea must be dif –
ferentiated from other diarrheal diseases of rabbits such
as coccidiosis, salmonellosis, Tyzzer disease, clostridial
enterotoxemia, enteric coronaviral enteritis, and colibac –
illosis. In weanling rabbits with severe disease, the pos –
sibility of dual infections should be considered, since
rotavirus infections are usually mild. Electron micros –
copy ( Ciarlet et al., 1998; Peeters et al., 1984; Schoeb et al.,
1986 ) or a capture enzyme-linked immunosorbent assay
can be used to detect rotavirus in feces ( Blutt et al., 2003;
Conner et al., 1988; De Leener et al., 2004; Hambraeus
et al., 1989; Thouless et al., 1986, 1988 ). A commercial
human rotavirus detection kit, utilizing immunochro –
matography, tested with fecal samples from rabbits with
diarrhea was shown to be applicable, as positive results
were confirmed by reverse transcription-polymerase
chain reaction (RT-PCR) and restriction endonuclease
analysis ( Fushuku and Fukuda, 2006 ). The electrophero –
type of rotaviruses can be determined by extraction of
nucleic acid from feces and electrophoresis in polyacryl –
amide gel ( De Leener et al., 2004; Herring et al., 1982 ).
Cytoplasmic fluorescence can be observed at the tips
of ileal villi using indirect immunofluorescence ( Petric
et al., 1978 ). Reverse transcription-polymerase chain
reaction (RT-PCR), immunohistochemistry, and in situ
hybridization can be used to detect rotavirus in formalin-
fixed tissues ( Tatti et al., 2002 ). Serum antibodies to rota –
virus can be detected by enzyme immunoassay ( Conner
et al., 1991; Hambraeus et al., 1989; Kelkar et al., 2004;
Thouless et al., 1988 ). Rabbits with rotaviral diarrhea
usually have no or low concentrations of antibody to
rotavirus, with a subsequent rise 2–4 weeks after onset
of signs ( Bryden et al., 1976; Sato et al., 1982 ). Unaffected
littermates develop high antibody concentrations, reflect –
ing concurrent subclinical infections.
CONTROL
Rotavirus is highly infectious and is transmitted by
fecal–oral spread; however, fomites cannot be excluded.
Transplacental transmission has not been demon –
strated. Infection appears to be acute and self-limit –
ing. Experimentally, virus is shed in the feces for about

389 RNA VIRUS INFECTIONS
II. RABBITS1 week following inoculation ( Castrucci et al., 1984 ;
Conner et at., 1988; Hambraeus et al., 1989; Petric et al.,
1978; Thouless et al., 1988 ). Recovered rabbits are refrac –
tory to challenge with homologous ( Castrucci et al., 1984;
Ciarlet et al., 1998; Conner et al., 1988, 1991; Hambraeus
et al., 1989 ) or heterologous rotaviruses ( Hambraeus et
al., 1989 ). Hence, cessation of breeding or quarantine of
the colony for 4–6 weeks, to prevent the introduction of
susceptible rabbits, should permit the infection to run
its course. As seropositive dams are not infectious, their
offspring should remain free of infection after maternal
antibodies disappear. Early weaning of rabbits with high
concentrations of passively acquired maternal antibodies
and removal to isolated facilities offer the possibility of
rederiving rabbits free of infection. Prevention of rotavi –
rus infection depends on barrier maintenance of rabbits.
Coronavirus Infections
Pleural Effusion Disease/Infectious
Cardiomyopathy Virus
HISTORY
During the 1960s in Scandinavia, increased mortal –
ity in European rabbits ( Oryctolagus cuniculus ) inoculated
with the Nichols strain of T reponema pallidum (Jorgensen,
1968 ) was attributed to a virus, named the Stockholm
agent, present in rabbit testicular emulsions ( Gudjonsson
et al., 1970 ). As the principal necropsy finding was pleu –
ral effusion, the name pleural effusion disease was sug –
gested ( Fennestad et al., 1975 ). Later, it was shown that
the major target organ was the heart, and the name infec –
tious cardiomyopathy was suggested, since the etiologic
agent appeared to be a coronavirus ( Small et al., 1979 ). The
resemblance of the clinicopathological features to feline
infectious peritonitis, a systemic coronavirus infection
of cats, was noted ( Osterhaus et al., 1982 ). The failure to
propagate the agent in vitro has precluded more definitive
characterization. It is unclear whether the agent is a natu –
rally occurring pathogen of rabbits or a virus from another
species adapted to rabbits in contaminated treponemal
stocks. However, the disease may be useful as a model
for virus-induced cardiomyopathy ( Alexander et al., 1992,
1993; Baric et al., 1990; Edwards et al., 1992 ).
ETIOLOGY
The disease, initially described in 1968, occurred in
rabbits inoculated with suspensions of rabbit testes con –
taining the Nichols strain of T. pallidum (Gudjonsson
and Skog, 1970; Jorgensen, 1968 ). Rabbits devel –
oped fever, circulatory insufficiency with pulmonary
edema, and pleural effusion and had high mortality
(Gudjonsson and Skog, 1970; Gudjonsson et al., 1970 ).
Subsequent inoculation of rabbits with pulmonary tis –
sue and pleural fluid from dead rabbits, previously inoculated with treponemes, resulted in a similar syn –
drome ( Gudjonsson et al., 1970 ). Differential filtration
of pleural fluid and serum from rabbits revealed an
infectious particle size of 25–50 nm, which is ether-sen –
sitive and inactivated at temperatures of 65°C or above
but not at 56°C ( Fennestad and MacNaughton, 1983;
Gudjonsson et al., 1972; Small et al., 1979 ). Sera from
rabbits contained pleomorphic virus particles, round
or elliptic, 75–100 nm in diameter, bearing club-shaped
projections 15–20 nm long ( Small et al., 1979 ), whereas
plasma from rabbits contained high concentrations of
pleomorphic virus-like particles measuring 51–98 nm
in diameter with projections 8–13 nm long ( Osterhaus
et al., 1982 ). Infectious sera, passed in various cell lines,
produced a cytopathic effect in primary rabbit kidney
and newborn human intestine cells ( Small et al., 1979 ).
However, the cellular pathogenic effect was lost after
two passages. Disease was not induced in mice, ham –
sters, or guinea pigs ( Gudjonsson et al., 1970; Small
et al., 1979 ).
In a complement fixation test, antigens in infectious
sera cross-reacted with antisera to the 229E (two-way
cross) and OC43 (one-way cross) strains of human coro –
navirus ( Small et al., 1979 ). Antisera to the rabbit car –
diomyopathy agent cross-reacted with feline infectious
peritonitis virus (FIPV), canine coronavirus (CCV), and
porcine transmissible gastroenteritis virus (TGEV) by
radioimmunoassay ( Small and Woods, 1987 ). Prior incu –
bation of the agent with antisera to CCV , FIPV , TGEV ,
or 229E virus reduced mortality in rabbits following
inoculation. However, prior immunization of rabbits
with CCV , FIV , or TGEV had little effect on survival.
Immunofluorescent staining of cardiac tissue from dis –
eased rabbits with antisera to 229E virus revealed antigen
in myocardial interstitial tissue, and rabbits surviving
infection developed antibodies to the virus in a comple –
ment fixation assay ( Small et al., 1979 ). In another study ,
however, surviving rabbits failed to demonstrate antibod –
ies against 229E virus in an enzyme immunoassay and
neutralization test, and against OC43 virus in a comple –
ment fixation test ( Fennestad and MacNaughton, 1983 ).
EPIDEMIOLOGY
When the disease was recognized in 1961, rab –
bits used in the serial propagation of T. pallidum had
a mortality rate of 2%. By the end of the 1960s, how –
ever, mortality had increased to 35–40% owing to con –
tamination of treponemal stocks with virus ( Fennestad,
1985; Jorgensen, 1968 ). There is no difference in disease
among breeds (Gudjonsson and Skog, 1970) or between
rabbits in the United States and Sweden ( Gudjonsson
et al., 1970 ). The agent was detected in T. pallidum –
infected rabbit tissues from Europe, the United States,
and Japan ( Fennestad et al., 1980 ). Nine isolates, obtained
from treponema-infected rabbits in various countries,

II. RABBITS
14. VIRAL DISEASES
390
exhibited a wide range in pathogenicity when inoculated
into rabbits ( Fennestad et al., 1986 ). Whereas virtually all
rabbits had fever, mortality ranged from 0 to 88%, and all
surviving rabbits resisted challenge with a virulent strain.
Two of four dams from litters inoculated at 6–9 days of
age became infected, whereas uninoculated littermates
or introduced cagemates failed to develop infection after
exposure for several months ( Fennestad et al., 1981 ).
Hence, transmission by direct contact occurs rarely .
CLINICAL SIGNS
Whereas clinical features vary by strain and pas –
sage of the agent, inoculated rabbits generally develop
a fever in 1–4 days which persists for 5–10 days. During
this early phase, acute deaths can occur without clini –
cal signs of illness ( Alexander et al., 1999 ). Clinical signs
are often consistent with those of congestive heart failure
including anorexia, weight loss, atony , tachypnea, and
dyspnea. Iridocyclitis has been reported. With virulent
strains, a third of infected rabbits die 3–5 days after inoc –
ulation, although deaths can occur until 14 days ( Baric et
al., 1990; Fennestad, 1985; Fennestad and MacNaughton,
1983; Fennestad et al., 1975, 1986; Fledelius et al., 1978;
Gudjónsson and Skog, 1970; Gudjonsson et al., 1970;
Small et al., 1979 ). Generalized or hindquarter muscu –
lar weakness was also reported in rabbits surviving the
acute phase of infection ( Gudjonsson et al., 1970; Small
et al., 1979 ). During the acute phase of infection, rab –
bits have a transient lymphopenia, followed by hetero –
philia ( Fennestad et al., 1975 ). Red cell indices were also
reduced but returned to normal by 6 weeks after inocula –
tion. There was a transient hypoalbuminemia; however
γ-globulin increased significantly ( Fennestad et al., 1975 ).
Serum potassium and lactate dehydrogenase were ele –
vated transiently . ECG abnormalities have been reported
during the acute phase (days 3–5) including tachycardia
in 74% of the infected rabbits, depressed R wave volt –
ages (16%), reduced T wave voltages (45%) and QT (cor –
rected) prolongation (32–42%) and correlated positively
with mortality ( Alexander et al., 1995, 1999 ). Ninety per –
cent of rabbits had persistent tachycardia in the subacute
phase (days 6–12 after infection), ECG changes persisted,
and echocardiographic changes in the subacute phase cor –
related positively with mortality . Surviving rabbits usually
returned to normal in 3–4 weeks. However, ECG abnor –
malities persisted after day 30 in some rabbits, reflecting
the development of dilated cardiomyopathy ( Alexander et
al., 1999 ). Reinoculation of surviving rabbits was without
effect, indicating development of resistance to the agent
(Fennestad, 1985; Fennestad and MacNaughton, 1983;
Fennestad et al., 1986; Gudjonsson et al., 1970 ).
PATHOLOGY
Rabbits dying during the acute phase of disease
have pulmonary edema, pleural effusion, and dilation of the right ventricle ( Alexander et al., 1992, 1993; Baric
et al., 1990; Christensen et al., 1978; Edwards et al., 1992;
Fennestad et al., 1975; Small et al., 1979 ). The pleural cav –
ities contain 2–50 ml of fluid, with or without fibrin, and
few cells; ascites may also occur in rabbits dying after the
first week. There are subepicardial and subendocardial
hemorrhages. Other findings may include hepatospleno –
megaly and congested lymph nodes. The heart weight
is increased, and there is dilation of the right ventricle
followed by dilation of the left ventricle. The cause of
death appears to be congestive heart failure ( Alexander
et al., 1992, 1993; Baric et al., 1990; Edwards et al., 1992;
Small et al., 1979 ). In fatal cases, there is lymphoid deple –
tion of the splenic follicles, focal degenerative changes
of the thymus and lymph nodes, and mild prolifera –
tive changes of renal glomeruli ( Christensen et al., 1978;
Small et al., 1979 ). In rabbits that survive the acute phase,
there is multifocal to diffuse myocardial degeneration
and necrosis, focal hepatic necrosis, and proliferative
changes in the spleen, lymph nodes, interstitial pulmo –
nary tissue, and renal glomeruli. Lesions similar to those
in the heart can occur in the diaphragm ( Small et al.,
1979 ). A mild non-suppurative, non-granulomatous ante –
rior uveitis develops during the acute phase of infection
and regresses within 4 weeks ( Fledelius et al., 1978 ). At
65 days after infection, myocarditis is evident, and about
one-third of rabbits had evidence of right- and/or left-
sided cardiac dilation ( Baric et al. 1990 ). After 2 years,
rabbits had pulmonary lymphoid hyperplasia, lymphoid
hyperplasia of lymph nodes and spleen, siderosis in the
spleen, necrosis and periportal inflammation in the liver
and interstitial fibrosis of the myocardium ( Fennestad et
al., 1986 ). No evidence of circulating or tissue immune
complexes was found ( Fennestad et al., 1981, 1986 ).
DIAGNOSIS
It remains to be determined whether the agent is a nat –
urally occurring pathogen of rabbits or perhaps a human
virus adapted to rabbits in contaminated stocks of the
Nichols strain of T. pallidum . The syndrome has only been
reported in experimentally inoculated rabbits. The clini –
cal course is characterized by onset of fever 1–3 days after
inoculation. Deaths may occur from 2–17 days after inocu –
lation, and mortality may exceed 75%. Other major signs
include ocular disease, anorexia, weight loss, tachypnea,
and atony . The gross and histological lesions are highly
characteristic of the disease. Antibodies to the virus cross-
react with the human coronavirus 229E and other mem –
bers of the group I mammalian coronaviruses (FlPV , CCV ,
and TGEV) ( Small and Woods, 1987; Small et al., 1979 ).
CONTROL
Because continued serial propagation of T. pallidum in
rabbits coincided with emergence of disease associated
with this agent, the possibility of contaminated stocks

391 RNA VIRUS INFECTIONS
II. RABBITSshould be considered ( Fennestad, 1985; Fennestad et al.,
1975; Gudjonsson et al., 1970 ). Rabbits inoculated with
virus-contaminated T. pallidum stocks that elicited no or
mild disease without mortality were protected from dis –
ease when challenged with stocks contaminated with
more pathogenic virus ( Fennestad, 1985; Fennestad
and MacNaughton, 1983; Fennestad et al., 1980, 1986;
Gudjonsson et al., 1970 ). This indicated that some T. pal –
lidum stocks were contaminated with a non-pathogenic
variant of the virus, which was able to confer protection.
Furthermore, isolates decreased in pathogenicity, as a
function of time, in rabbits that survived acute infection
(Fennestad, 1985; Fennestad et al., 1986 ). Convalescent
sera from rabbits that survived infection partially pro –
tected challenged rabbits against mortality but not dis –
ease ( Fennestad et al., 1981; Gudjonsson et al., 1970;
Small et al., 1979 ). Passage of contaminated T. pallidum
stocks through hamsters removed the agent responsible
for disease ( Fennestad et al., 1980; Skovgaard Jensen,
1971 ). Subsequent reintroduction of these T. pallidum
stocks in rabbits was without effect. Similarly, inocula –
tion of rabbits, with popliteal lymph nodes from surviv –
ing rabbits, transferred treponemes but not the agent
(Gudjonsson et al., 1970 ).
Rabbit Enteric Coronavirus
HISTORY
A coronavirus was detected in the feces of rabbits with
diarrhea by LaPierre et al. (1980) in Canada. Rabbits were
6–10 weeks old and from several colonies. Subsequently,
coronavirus was detected in young rabbits with diarrhea
in several European countries ( Eaton, 1984; Osterhaus
et al., 1982; Peeters et al., 1984 ). Although the virus
appears to be readily detected in rabbits by workers in
Canada ( Descoteaux et al., 1985; LaPierre et al., 1980 ),
the failure to propagate the agent in vitro has ham –
pered investigations of its prevalence in European rabbit
(Oryctolagus cuniculus ) populations.
ETIOLOGY
Feces of rabbits with diarrhea exhibit hemagglutina –
tion activity with rabbit erythrocytes, particularly with
a peak of 1.18 g/cm3 from a sucrose density gradient
(Descoteaux et al., 1985; LaPierre et al., 1980 ). Electron
microscopy of fecal samples revealed pleomorphic par –
ticles with an inner diameter of 60–220 nm and surface
projections of 20 nm ( Eaton, 1984; LaPierre et al., 1980 ).
Particles obtained from a Percoll gradient appeared as
spherical enveloped particles 40–50 nm in size with pro –
jections of 10–12 nm ( Descoteaux et al., 1985 ). In rabbits
inoculated orally, pleomorphic particles 60–90 nm, with
surface projections of 10 nm, were detected in the feces
(Osterhaus et al., 1982 ). Electron microscopy of fecal
samples using antisera to rabbit coronavirus revealed immune aggregates of viral particles with morphologi –
cal features of coronaviruses ( Descoteaux et al., 1985 ).
Analysis of structural polypeptides by immunoblot –
ting revealed that antisera to avian infectious bronchitis
virus (AIBV) and TGEV detected many of the same epi –
topes as antisera to the rabbit coronavirus ( Descoteaux
et al., 1985 ). Immune sera to 229E, but not AIBV nor
porcine hemagglutinating encephalitis virus, inhibited
the hemagglutination activity of the rabbit coronavi –
rus. No cytopathic effect occurred following inoculation
of various cell lines with fecal samples ( Eaton, 1984;
LaPierre et al., 1980 ).
EPIDEMIOLOGY
In a survey of 130 diarrheic rabbits from 21 rabbit –
ries, coronavirus was detected in the cecal contents of
one rabbit in association with Escherichia coli infection
(Peeters et al., 1984 ). Rabbits with diarrheal disease
associated with coronavirus infection are usually 3–10
weeks old ( Eaton, 1984 ; La Pierre et al ., 1980). Virus has
also been detected in the feces of apparently healthy
rabbits ( Descoteaux et al., 1985 ). Experimentally,
orally inoculated rabbits shed virus for up to 29 days
(Descoteaux and Lussier, 1990 ). A serological sur –
vey of 238 Oryctolagus cuniculus from six rabbitries in
Washington state revealed that 23 (10%) had antibodies
to canine coronavirus ( Deeb et al., 1993 ).
CLINICAL SIGNS
In an outbreak of enteric disease in 3–8-week-old
rabbits in a barrier-maintained breeding colony, rab –
bits exhibited lethargy, diarrhea, and swollen abdo –
mens ( Eaton, 1984 ). Between 40 and 60% of rabbits were
affected, and virtually all died within 24 h after onset of
signs. LaPierre et al. (1980) also reported rapid death
following the onset of clinical signs in the majority of
affected rabbits. Experimentally, rabbits developed soft
feces ( Osterhaus et al., 1982 ) and transient watery diar –
rhea ( Descoteaux and Lussier, 1990 ) after inoculation.
None of the rabbits died ( Descoteaux and Lussier, 1990 ).
PATHOLOGY
In rabbits with diarrhea, the perianal region is soiled
with feces ( Eaton, 1984 ). Rabbits appear cachectic and
dehydrated. Although the stomach and intestines are
unaffected, the cecum is distended with watery fluid
(Eaton, 1984; Peeters et al., 1984 ). Histologically, in the
small and large intestines there is a diffuse infiltration
of inflammatory cells and mucosal edema ( Eaton, 1984 ).
In an experimental study, the small intestines were con –
gested and the cecal contents watery ( Descoteaux and
Lussier, 1990 ). Microscopically, at 6 hours after inocu –
lation, there was necrosis of enterocytes at the tips of
intestinal villi. Villous atrophy and hypertrophic crypts

II. RABBITS
14. VIRAL DISEASES
392
were present 2–3 days after inoculation. However, by 6
days, microscopic lesions were absent.
DIAGNOSIS
Rabbits with diarrheal disease are usually recently
weaned and 3–10 weeks old ( Eaton, 1984; LaPierre
et al., 1980 ). Because the rabbit coronavirus agglutinates
erythrocytes, the hemagglutination activity of the feces
provides an indication of the presence of virus ( LaPierre
et al., 1980 ). Coronavirus particles can be demonstrated
in the feces by electron ( Eaton, 1984; LaPierre et al., 1980;
Osterhaus et al., 1982; Peeters et al., 1984 ) and immuno –
electron ( Descoteaux and Lussier, 1990; Descoteaux et al.,
1985 ) microscopy. The latter technique is more sensitive
in detecting virus particles ( Descoteaux et al., 1985 ). At
necropsy, the cecal contents are fluid ( Descoteaux and
Lussier, 1990; Eaton, 1984 ); histologically, there is intes –
tinal villous atrophy ( Descoteaux and Lussier, 1990 ).
Adult rabbits may have antibodies in the hemaglutina –
tion inhibition assay ( LaPierre et al., 1980 ). As experi –
mental inoculation of rabbits with coronavirus failed to
mimic the field disease with its high morbidity and mor –
tality, consideration should be given to the existence of
co-pathogens ( Descoteaux and Lussier, 1990; Osterhaus
et al., 1982 ). In two studies where other agents were
considered, coronavirus was associated with Esherichia
coli and Clostridium perfringens infections ( Eaton, 1984;
Peeters, et al., 1984 ).
CONTROL
Because only one naturally occurring outbreak of
diarrheal disease associated with coronavirus has been
reported ( Eaton, 1984 ), there is scant information on con –
trol of the disease. The feeding of hay, delay in weaning,
and administration of coccidiostats and antibiotics were
ineffective in preventing mortality.
Calicivirus Infections
The etiologic agents of necrotic hepatitis of leporids
have profoundly affected world rabbit populations
since 1984. They are classified as members of the fam –
ily, Caliciviridae , and the genus, Lagovirus . This genus
contains two distinct species which cause clinical dis –
ease denoted as Rabbit hemorrhagic disease virus (RHDV)
and European brown hare syndrome virus (EBHSV) ( Green
et al., 2000 ). These diseases were first recognized in the
mid-1980s. RHD was reported in China in 1984 and
subsequently in other countries. EBHS was observed
in Europe several years before RHD was diagnosed in
domestic European rabbits. Antibodies to RHDV were
in archived serum from 1961 in the Czech Republic
and Austria ( Nowotny et al., 1997 ) and viral RNA in
serum samples from Britain stored in 1955 ( Moss et al., 2002 ). Antibodies to EBHSV were found as early as 1962
(Frölich and Lavazza, 2008 ) in serum in England and
viral RNA in tissues collected in the 1970s in Sweden
(Bascunana et al., 1997 ).
A third member of the genus Lagovirus was identi –
fied in domestic rabbits in Italy in 1996 and was named
rabbit calicivirus (RCV) ( Capucci et al., 1996 ). No clini –
cal signs were associated with RCV infection although
seroconversion resulted in protection from RHDV
(Capucci et al., 1997 ). A closely related strain, denoted
rabbit calicivirus, Australia 1 (RCV-A1) was recently
isolated from wild European rabbits in Australia and
partially sequenced ( Strive et al., 2009 ). Whether anti –
bodies which recognize RHDV epitopes in retrospective
studies are associated with RCV and its variants rather
than RHDV is unknown. Another closely related virus,
denoted Michigan Rabbit Calicivirus (MRCV) was iso –
lated from an outbreak of hemorrhagic disease in the
United States in 2001 (Bergin et al., 2009). One mem –
ber of the genus Vesivirus , family Caliciviridae has been
described as a potential intestinal pathogen of rabbits
(Martín-Alonso et al., 2005 ).
Rabbit Hemorrhagic Disease Virus (RHDV)
HISTORY
In early 1984, an acute fatal disease of rabbits was
reported from many regions of China ( Xu and Chen,
1989 ; Xu et al ., 1988). The disease was unlike any pre –
viously reported syndrome and may have originated
in rabbits imported from Europe. Tests of rabbit sera
stored in Czechoslovakia in 1978 revealed antibodies to
the virus ( Rodak et al., 1990a ). Although the syndrome
was initially variously named, it came to be known as
RHD. Subsequently, the disease was reported in sev –
eral European countries in 1987 and 1988 ( Gregg and
House, 1989; Parra and Prieto, 1990; Patton, 1989 ). The
negative impact of this disease on wild rabbit popu –
lations in Europe has been significant ( Calvete et al.,
2002; Marchandeau et al., 1998 ). Currently, it is endemic
in most of Europe, Asia, Africa, Australia, and New
Zealand.
By late 1988, the virus reached North America and
was reported from many locations in Mexico ( Gregg
and House, 1989; Gutierrez, 1990; Patton, 1989 ). Mexico
mounted an eradication campaign and has since
remained free of the virus ( McIntosh et al., 2007 ). Four
outbreaks of RHDV were reported in the United States
between 2000 and 2007 ( Anonymous, 2000; Campagnolo
et al., 2003; U.S.D.A., 2005; McIntosh et al., 2007 ). Each
time the virus was contained, but how it was introduced
was not determined ( McIntosh et al., 2007 ).
In 1995, a laboratory strain of RHDV was being
tested in field experiments under quarantine conditions
at Wardang Island, Australia, to evaluate the virulence,

393 RNA VIRUS INFECTIONS
II. RABBITStransmissibility, and persistence of RHDV ( McColl
et al., 2002 ). It was also being considered as a potential
biological control agent for rabbits. The virus escaped
and spread, first to other rabbits on Wardang Island,
and then to the mainland. It resulted in significant
mortality immediately, reducing rabbit populations
in some areas by 95% ( Mutze et al., 1998 ). It is cur –
rently endemic throughout Australia. In 1996, the New
Zealand government was asked to consider the use of
RHDV as a biological control agent ( Thompson and
Clark, 1997 ). The government denied these requests;
however, RHDV was introduced to New Zealand ille –
gally by farmers hoping to reduce local rabbit popula –
tions and is now widely distributed in both the North
and South Islands of New Zealand ( O’Keefe et al., 1999;
Thompson and Clark, 1997 ).
ETIOLOGY
A virus has been consistently purified from the liver
and spleen of affected rabbits, the characteristics of
which have been described by several investigators
(Du, 1990, 1991; Du et al., 1986; Liebermann et al., 1992 ;
Ohlinger and Theil, 1991; Sihid et al., 1989; Smid et al.,
1989; Xu, 1991; Xu and Chen, 1989 ; Xu et al., 1988). The
virus is non-enveloped, spherical, and 28–34 nm in diam –
eter and has a buoyant density of 1.32–1.38 g/cm3. The
icosahedral capsid has 32 cylindrical capsomeres, 5–6 nm
in diameter, comprised of four structural polypeptides.
The virus is a member of the family, Caliciviridae , and is
the type virus of the genus, Lagovirus (Green et al., 2000 ).
RHDV was completely sequenced and has a positive
strand RNA genome of 7437 kb ( Meyers et al., 1991a ).
The genomic organization is similar to EBHSV with
two open reading frames (ORF) rather than the three
ORFs found in other caliciviruses ( Wirblich et al., 1996 ).
The first ORF encodes non-structural proteins as well
as the large capsid protein and the second ORF encodes
an additional capsid protein. One subgenomic RNA
(2.2 kb) is present ( Ohlinger et al., 1990; Parra and Prieto,
1990 ). Amino acid identity between RHDV and EBHSV
is approximately 76% ( Wirblich et al., 1994 ). The virus
agglutinates erythrocytes of humans and guinea pigs,
but not rabbits. Non-hemagglutinating strains of RHDV
have been identified ( Tian et al., 2007 ). While the major –
ity of strains of RHDV have a high level of amino acid
identity, antigenic variants have been described ( Capucci
et al., 1998; Asgari et al., 1999; Shirrmaier et al., 1999 ).
The virus is highly stable in the environment and
sufficient infectious RNA is present to infect rabbits
from liver samples stored for 20 days at 22°C ( McColl
et al., 2002 ). In a separate study, infectious virus was
injected into bovine liver and then the liver was
exposed outdoors to ambient temperatures (2–34°C)
(Henning et al., 2005 ). Sufficient viral particles were present at the longest time point studied, 91 days, to
infect rabbits when inoculated undiluted. Viral infectiv –
ity is unaffected by treatment with ether, chloroform,
exposure to pH 3, or heating at 50°C. However, the
virus is inactivated by 1% sodium hydroxide or 0.4%
formaldehyde at ambient temperature, 4°C or 37°C. The
virus fails to grow in many primary and established cell
lines but has been adapted to a transformed rabbit kid –
ney cell line ( Ji et al., 1991; Xu, 1991 ). Rabbits may be
the only species susceptible to infection, as inoculation
of guinea pigs, hamsters, rats, or mice failed to induce
disease. Experimentally, inoculation of European hares
(Lepus europaeus ) failed to induce disease ( Lavazza et al.,
1996; Smid et al., 1991 ).
EPIDEMIOLOGY
Since the report of the first outbreak of RHDV in
China in 1984, RHDV has rapidly spread through –
out the world. The disease has been reported from
China, Korea, most European countries, Asia, Africa,
North America, Australia. and New Zealand (Chasey
et al., 1997; McIntosh et al., 2007; Morisse et al., 1991 ).
Countries experiencing epidemics with high morbidity
and mortality include China ( Xu and Chen, 1989 ), Korea
(Lee and Park, 1987 ), Italy ( Cancellotti and Renzi, 1991;
Patton, 1989 ), Spain ( Parra and Prieto, 1990; Villafuerte
et al., 1994 ); France ( Morisse et al., 1991 ), Germany
(Loliger and Eskens, 1991 ), Mexico ( Gregg and House,
1989; Gutierrez, 1990 ), Taiwan ( Shien et al., 1998 ), Cuba
(Farnós et al., 2007 ), Saudi Arabia ( Abu Elzein and
Al-Afaleqand, 1999 ), Bahrain ( Forrester et al., 2006 ),
Tunisia ( Bouslama et al., 1996 ), the United Kingdom
(White et al., 2004 ), and the United States ( McIntosh et
al., 2007 ). The disease has also been reported from India
(Sundaram et al., 1991 ) and the Middle East ( Kuttin et
al., 1991 ). Viruses from China, Korea, and Europe appear
to be similar ( Du, 1990; Gregg et al., 1991 ), although
evolution of viruses in restricted geographic areas has
occurred ( Muller et al., 2009 ). Recently, a virus isolated in
the Netherlands was demonstrated to be 99% identical to
RHDV from France ( van de Bildt, et al., 2006 ).
There have been four outbreaks of RHDV in the US; in
Iowa in 2000 ( Anonymous, 2000 ), in Utah in 2001, which
was transferred to Illinois by shipment of infected rabbits
(Campagnolo et al., 2003 ), in a zoo in New York in 2001
(McIntosh et al., 2007 ), and in Indiana in 2005 ( McIntosh
et al., 2007 ; U.S.D.A., 2005). Isolates from these outbreaks
were shown by comparative genomic analysis to be
closely related to isolates from China, but to have sepa –
rate origins ( McIntosh et al., 2007 ).
There is evidence that RHDV was present in wild and
domesticated populations of rabbits in Europe ( Moss
et al., 2002; Rodak et al., 1990a ), Australia ( Cooke et al.,
2000; Nagesha et al., 2000 ), and New Zealand ( O’Keefe
et al., 1999 ) prior to the recognition of RHDV . Studies in

II. RABBITS
14. VIRAL DISEASES
394
England demonstrated serological evidence as well as
partial RNA sequences recovered from tissues or serum
of apparently healthy rabbits ( Moss et al., 2002; White
et al., 2004 ). These partial sequences were more closely
related to RHDV than RCV , suggesting that non-patho –
genic RHDV viruses have circulated in parts of the world
for some time.
No apparent difference in susceptibility to infec –
tion among breeds of Oryctolagus cuniculus has been
reported ( Xu and Chen, 1989 ; Xu et al ., 1988). Outbreaks
of disease have occurred mainly in rabbits 2 months of
age and older, whereas younger rabbits were clinically
unaffected. Of the rabbits that survived the infection,
viral RNA has been detected in various organs includ –
ing the spleen, liver, mesenteric lymph node, and bile
for 15 weeks after infection suggesting that a chronic
carrier state can occur ( Gall et al., 2007 ).
Transmission of virus is horizontal, primarily by direct
contact with secretions and excretions of infected rabbits
and fecal–oral spread may be the major mode of transmis –
sion. Contaminated fomites, such as feed, water, utensils,
and animal attendants, may be important in transmission
as well. Transmission by insects has been suggested as an
explanation for the spread of RHDV from Wardang Island
to the mainland. Laboratory experiments demonstrated
that RHDV can be transmitted by blowflies ( Phormia sp.),
bushflies ( Musca vetustissima ) and mosquitoes ( Gould
et al., 1997; McColl et al., 2002 ). Transmission by the lesser
brown blowfly , Calliphora dubia was suggested by evi –
dence that RHDV RNA was detected in fly feces after flies
were fed a meal from a RHDV-infected liver ( Asgari et al.,
1998 ). There is no evidence of vertical transmission. Fecal
shedding by carnivores after ingestion of RHDV-infected
carcasses has been suggested as a potential mode of
spread. Serological evidence of infection in foxes has been
demonstrated in several studies, although no infectious
particles have been demonstrated ( Frölich et al., 1998;
Leighton et al., 1995 ; Philbey et al ., 2005). Experimentally ,
the routes of entry , in order of importance, are oral, con –
junctival, nasal, and skin trauma. In China and Europe,
epidemics of disease usually begin in November and end
in March. Although Oryctolagus cuniculus appears to be
the only species susceptible to disease, hares in China also
appear to be infected and capable of transmitting the virus
to rabbits experimentally ( Xu, 1991 ).
A serological survey of 1461 rabbits from 43 farms,
apparently free of viral hemorrhagic disease in
Czechoslovakia revealed that 283 rabbits (19%) from 33
farms had anti-RHDV antibodies ( Rodak et al., 1990a ).
Using another set of sera, collected between 1975 and
1987 from laboratory-maintained rabbits, antibodies
were detected in 32 of 42 sera (76%). This suggests that
rabbit colonies in Czechoslovakia harbored a viral agent
with characteristics similar to those of Rabbit hemor –
rhagic disease virus, but with lower pathogenicity, several years before the syndrome was recognized in
China.
RHDV has been shown to bind to histo-blood group
antigens expressed on the surface of respiratory and
digestive tract epithelial cells ( Ruvoën-Clouet et al.,
2000 ). This binding may partially explain the differential
susceptibility of young versus older rabbits ( Rademacher
et al, 2008; Ruvoën-Clouet et al., 2000 ). In addition, poly –
morphisms of these antigens in wild rabbits in France
have been linked to differential survival from RHDV
infection ( Guillon et al., 2009 ).
CLINICAL SIGNS
In general, the disease is acute and highly infectious,
with high morbidity and mortality ( Marcato et al.,
1991; Xu and Chen, 1989 ; Xu et al., 1988). The incuba –
tion period ranges from 1–2 days. Morbidity is 70–80%,
and mortality approaches 100%. During outbreaks, the
number of rabbits affected usually peaks in 2–3 days
and lasts 7–13 days. After viral introduction, rabbits die
suddenly with few clinical signs. Rabbits become febrile
and exhibit depression, lethargy, and anorexia. Other
clinical signs include tachypnea, cyanosis, abdominal
distention, and constipation or diarrhea. Because the
disease is acute, clinical signs may be brief and often are
unnoticed. In the terminal stage, rabbits become hypo –
thermic, recumbent, and have convulsions and epi –
staxis. Surviving rabbits exhibit depression, anorexia,
and fever which usually abates in 2–3 days. In endemic
areas, the form of disease observed in surviving rab –
bits is more common. Hematologic evaluation usually
reveals a lymphopenia, a gradual decline in thrombo –
cytes, and prolonged prothrombin and thrombin times.
A paracoagulation test with protamine sulfate gives a
strong positive reaction. Fibrin degradation products
can be detected in most moribund rabbits.
In young rabbits, clinical signs of infection are typi –
cally absent ( Ferreira et al., 2004, 2006; Mikami et al.,
1999 ). However, liver inflammation as evidenced by
increase in hepatic transaminases and increase in hetero –
phil liver infiltrate occurs, although there is little hepa –
tocyte damage ( Ferreira et al., 2004, 2006; Mikami et al.,
1999; Prieto et al., 2000 ) and less replication of the virus
within hepatocytes than adults as evidenced by immu –
nochemical staining (Preito et al., 2000). Elevation of
transaminases persists for at least 3 weeks after infection
(Ferreira et al., 2004 ). Young rabbits infected with RHDV
develop a protective antibody response that persists and
protects them from infection as adults ( Ferreira et al.,
2008 ). However, these rabbits may serve as healthy carri –
ers of the virus ( Ferreira et al., 2004, 2008 ).
PATHOLOGY
The pathological changes apparently result from vire –
mia, with death attributable to an acute disseminated

395 RNA VIRUS INFECTIONS
II. RABBITScoagulopathy with deep venous thrombosis ( Gregg and
House, 1989; Gregg et al., 1991; Marcato et al., 1991; Xu
and Chen, 1989; Xu et al., 1986 , 1988). Grossly, conges –
tion and hemorrhage occur in most organs but are most
pronounced in the lungs ( Figure 14.3 ). The liver is pale
and has a fine reticular pattern of periportal necrosis, the
most consistent finding in the disease ( Figure 14.4 ). The
spleen and kidneys may be dark and swollen owing to
acute infarction ( Figure 14.4 ). Lymph nodes are edema –
tous and may contain petechial hemorrhages. There is
often segmental catarrhal enteritis.
In most fatal cases, rabbits die from a severe and
massive intravascular coagulopathy ( Gregg et al., 1991 ).
The disease consistently causes acute hepatic necrosis,
which may be the only lesion found. Hepatic necrosis is periportal and diffuse, and when severe, may bridge
acini and cause dissociation of hepatic cords ( Gregg
et al., 1991; Percy and Barthold, 2007 ). Small single or
multiple intranuclear inclusion bodies can be found
in degenerate hepatocytes. There is often little inflam –
mation in necrotic areas. Many tissues, especially the
lungs, spleen, and kidneys, may have varying degrees
of congestion and hemorrhage due to microinfarction
or major venous thrombi. Acute coagulative necro –
sis due to microinfarction may be found in any organ.
Microinfarcts in the brain account for the terminal neu –
rological signs. Pulmonary venous thrombosis accounts
for the frothy serosanguinous discharge from the nares
seen terminally. A segmental necrotizing enteritis with
severe crypt necrosis and villous atrophy can be found
(Gregg et al., 1991 ). Lymphoid tissues may have vary –
ing degrees of degeneration and karyorrhexis of lym –
phocytes. The spleen and thymus are more often
affected than lymph nodes.
DIAGNOSIS
A presumptive diagnosis can be made on the basis
of the epidemiological features, clinical signs, and
pathological findings ( Xu and Chen, 1989 ). Virus can
be detected with RT-PCR using multiple tissues includ –
ing blood ( Gall et al., 2007; Guittré et al., 1995; Shien et
al., 2000; Vende et al., 1995; Wang et al., 2008; Yang et
al., 2008 ). No satisfactory in vitro isolation methods
have been described, and rabbit inoculation is the only
method available to isolate or propagate a new strain
of virus (O. I. E. 2008). Immunocapture RT-PCR was
developed to simplify the tissue processing required to
analyze numerous samples (Le Gall-Reculé et al., 2001).
By electron microscopy, viral particles, 28–34 nm in
diameter, are frequently found in hepatocytes ( Valicek
et al., 1990; Xu and Chen, 1989 ). The hemagglutina –
tion test, with type O human erythrocytes, is useful for
detection of virus in suspensions of liver, lungs, spleen,
and kidneys, from infected rabbits ( Shien et al., 2000;
Xu and Chen, 1989 ). Antisera or monoclonal antibodies
to the virus can detect virus in hepatocytes in infected
rabbits by immunoenzyme and immunofluorescence
tests ( Berninger and House, 1995; Capucci et al., 1991;
Carrasco et al., 1991; Gregg and House, 1989; Gregg et al.,
1991; Park and Itakura, 1992; Rodak et al., 1990b, 1991;
Stoerckle-Berger et al., 1992 ). Serological tests include
the hemagglutination inhibition test and the enzyme-
linked immunosorbent assay, the latter being preferable
because of increased sensitivity and specificity ( Chasey
et al., 1995; Cooke et al., 2000; Rodak et al., 1990b ). Both
indirect sandwich ELISAs and competitive-ELISAs have
been developed for detection of antibodies ( Guittré et al.,
1995 ). Seroconversion to RHDV without clinical signs
of disease is suggestive of RCV infection and should be
interpreted with caution ( Capucci et al., 1997 ).
FIGURE 14.3 Diffusely congested lungs from rabbit found dead
with RHD. Courtesy of Dr. D. Gregg.
FIGURE 14.4 Reticular pattern of liver (bottom, center) consistent
with periportal necrosis in a rabbit found dead with RHD. The kidney
(top, center) is diffusely congested. Courtesy of Dr. D. Gregg.

II. RABBITS
14. VIRAL DISEASES
396
CONTROL
Measures to prevent introduction of the virus include
restricted access and disinfection of all equipment entering
or leaving a facility ( Xu and Chen, 1989 ). Cages and equip –
ment can be disinfected with 0.5% sodium hypochlorite
or 1% formalin. Rabbits from endemic areas should not
be introduced directly into the colony , but quarantined
for at least 1 month and tested serologically for anti-
RHDV antibodies. Similarly , colonies with disease should
be quarantined and depopulated, since surviving rabbits
shed virus for at least a month and possibly longer ( Gregg
and House, 1989; Gregg et al., 1991 ). Antiserum has been
shown to be protective ( Du, 1990; Huang, 1991 ). Tissue-
derived vaccines, inactivated with formaldehyde ( Du,
1990; Du et al., 1986; Huang, 1991; Xu and Chen, 1989 ) or
β-propiolactone ( Arguello Villares, 1991; Smid et al., 1991 ),
have been shown to be safe and efficacious in prevent –
ing disease. Resistance develops within 1–2 weeks after
vaccination and lasts for 5–15 months. Although disease
may be prevented in vaccinated rabbits, persistent infec –
tion may develop on exposure to the virus ( House et al.,
1990 ). Thus, vaccinated rabbits infected with virus should
be considered infectious and should not be introduced
into previously unexposed rabbitries. Additional vaccines
have been developed, including those combining protec –
tion against both RHDV and myxomatosis ( Bertagnoli et
al., 1996; Calvete et al., 2004; Fernández-Fernández et al.,
2001; Torres et al., 2001 ). Since the decline of rabbit popu –
lations in southern France and the Iberian peninsula has
affected the survival of rabbit-prey species ( Marchandeau
et al,. 2000 ), vaccination of free-living rabbits has been
undertaken in a field trial on an island off Spain and sub –
sequently on the mainland ( Calvete et al., 2004; Torres et
al., 2001 ).
European Brown Hare Syndrome Virus (EBHSV)
HISTORY
A disease, characterized by hemorrhages in the trachea
and lungs, pulmonary edema, and necrotic hepatitis,
with high mortality, has been observed since 1980–1985
in European hares ( Lepus europaeus ) and Mountain hares
(Lepus timidus ) in many European countries ( Billinis
et al., 2005 ; Duff et al., 1994; Frölich 1996; Frölich et al.,
2001, 2003a, 2007; Gavier-Widen and Morner, 1991; Le
Gall-Reculé et al., 2006; Morisse et al., 1990, 1991; Syrjälä
et al., 2005 ). The virus has also been detected in European
hares translocated to Argentina ( Frölich et al., 2003b ). The
disease, named European brown hare syndrome (EBHS),
is similar to RHD, but affects hares.
ETIOLOGY
Electron microscopy of hepatocytes from affected
hares reveals non-enveloped, icosahedral particles about
30 m in diameter ( Chasey and Duff, 1990; Marcato et al., 1991 ). Immunoblotting of the virus reveals a major struc –
tural protein of about 60 kDa, similar to RHDV; however,
other proteins of the latter virus are absent, suggesting
that the viruses are similar but not identical ( Capucci
et al., 1991; Ohlinger and Thiel, 1991 ). Sequencing of the
genome ( Gould et al., 1997; Le Gall et al., 1996 ; Meyers
et al., 1991b; Nowotny et al., 1997; Parra et al., 1993;
Rasschaert et al., 1995; Wirblich et al., 1994 ) revealed that
RHDV and EBHSV have similar genomic organization
and are both members of the family, Caliciviridae , and the
genus, Lagovirus , however they represent two distinct
species ( Green et al., 2000 ). EBHSV and RHDV have been
shown to have approximately 76% amino acid identity
(Wirblich et al., 1994 ).
EBHSV fails to grow in primary hare and rabbit cell
lines ( Gavier-Widen and Morner, 1991; Henriksen et al.,
1989 ), and inoculation of mice and guinea pigs failed to
induce disease ( Henriksen et al., 1989 ). In general, inoc –
ulation of rabbits ( Oryctolagus cuniculus ) with EBHSV ,
or tissues from infected hares, has not resulted in clini –
cal disease in rabbits ( Capucci et al., 1991; Chasey et al.,
1992; Eskens and Volmer, 1989; Nauwynck et al., 1993 )
or cross-protective immunity ( Nauwynck et al., 1993 ),
although clinical disease similar to RHD was induced
by Morrise et al. (1990) following inoculation of liver
homogenates from affected hares in France.
EPIDEMIOLOGY
The disease has been reported in wild hares from sev –
eral European countries, including England ( Chasey and
Duff, 1990 ), the Czech Republic ( Nowotny et al., 1997 ),
Greece ( Billinis et al., 2005 ), Finland ( Syrjälä et al., 2005 ),
Slovakia ( Frölich et al., 2007 ), Switzerland ( Frölich et al.,
2001 ), France ( Le Gall-Reculé et al., 2006 ), Poland ( Frölich
et al., 1996, 2003a ), as well as South America ( Frölich
et al., 2003b ). EBHSV has also been reported from breed –
ing farms for hares in Denmark ( Henriksen et al., 1989 )
and Sweden ( Gavier-Widen and Morner, 1991 ). In France,
Germany , and Italy , the geographic distribution of EBHS
coincides with RHD in wild and domesticated rabbits
(Cancellotti and Renzi, 1991; Loliger and Eskens, 1991;
Morisse et al., 1991 ). Seroprevalence has been determined
in many countries. Thirty-eight percent of hares tested in
Poland were positive for EBHSV antibodies ( Frölich et al,
1996 ), 29% in Germany (Frölich et al., 2003) while 73% of
hares shot by hunters in Slovakia had anti-EBHSV anti –
bodies, suggesting some variation in the prevalence of
infection.
Similar to RHDV , young hares are resistant to the
virus ( Morisse et al., 1991 ). The major mode of trans –
mission appears to be fecal–oral ( Morisse et al., 1991 ).
Contamination of feed and water with excreta from
infected hares is probably a common mode of transmis –
sion ( Gavier-Widen and Morner, 1991 ). Among wild

397 RNA VIRUS INFECTIONS
II. RABBITSand farmed hares in Denmark and Sweden, mortality is
seasonal, beginning in October and continuing through
March ( Gavier-Widen and Morner, 1991; Henriksen
et al., 1989 ), presumably as there are more young sus –
ceptible hares at the end of the breeding season. Recent
evidence from Finland indicates that the majority of the
cases occurred in the spring and summer, with a minor
peak in October and November ( Syrjälä et al., 2005 ).
The hare population in Finland was lowest in winter
and this may influence transmission. In England, excess
mortality in wild hares was reported in September
through November ( Chasey and Duff, 1990 ).
CLINICAL SIGNS
In general, the disease is acute and highly infectious,
with high morbidity and mortality ( Henriksen et al.,
1989; Marcato et al., 1991 ). Clinical signs include depres –
sion, anorexia, muscular tremors, incoordination, paraly –
sis, convulsions, and occasionally epistaxis. Death occurs
5–24 h after onset of signs, and affected hares rarely
recover. The reported morbidity is 75%, and mortal –
ity approaches 100% ( Gavier-Widen and Morner, 1991;
Henriksen et al., 1989 ). In Argentina ( Frölich et al., 2003b )
and Slovakia ( Frölich et al., 2007 ) evidence of a chronic,
less pathogenic form of EBHSV has been demonstrated.
In Argentina, no clinical signs suggestive of EBHSV were
reported; however 11% of 80 spleen samples were posi –
tive for EBHSV antigen ( Frölich et al., 2003b ) whereas
antibodies to EBHSV were present in only one sample. In
Slovakia, antibodies against EBHSV were found in 73% of
86 sera tested although hare populations were reported
to be stable with no increase in mortality noted before or
during the sampling period ( Frölich et al., 2007 ).
PATHOLOGY
Death is attributable to multiple organ failure result –
ing in pulmonary edema and hemorrhage, adrenocor –
tical necrosis, renal circulatory disorders, and hepatic
necrosis ( Henriksen et al., 1989; Marcato et al., 1991 ).
Gross examination of hares reveals marked pulmonary
congestion and edema, as well as hepatic congestion and
hemorrhages. Moderate splenomegaly and gastric ulcer –
ation are detected in some hares. A catarrhal to necrotiz –
ing conjunctivitis may be present. Microscopically, there
is diffuse acute coagulation necrosis of hepatic periportal
and midzonal areas, accompanied by formation of acido –
philic bodies (GavierWiden and Morner, 1991; Henriksen
et al., 1989 ). In many hares, there is basophilic stippling
in the cytoplasm of hepatocytes in periportal areas, rep –
resenting granular calcification. Many livers have micro –
vacuolar fatty degeneration. About one-fourth of affected
hares have splenic cellular depletion and hyaline-like
changes in the sinuses and cords, and about one-third
have renal tubular necrosis and calcification. In the brain,
cerebral neurons and cerebellar Purkinje cells exhibit granulovacuolar degeneration. In apparently healthy
hares with antibodies to the virus, hepatic vacuolar
degeneration, tracheitis, and hyperplasia of splenic fol –
licles are observed ( Marcato et al., 1991 ).
DIAGNOSIS
Diagnosis is similar to that for RHD. RT-PCR has
been used to detect virus in tissues ( Bascunana et al.,
1997; Gould et al., 1997 ). The hemagglutination test
for detection of virus in tissue specimens is less sensi –
tive ( Capucci et al., 1991 ). Use of two ELISAs, in series,
employing monoclonal antibodies, in which the first
test detects group antigen and the second virus-specific
antigen, has also been used to detect virus ( Capucci et
al., 1991 ). The ELISA is preferred for detection of anti –
bodies ( Capucci et al., 1991 ).
CONTROL
Methods of control in breeding farms for hares are
similar to those for rabbit hemorrhagic disease. Preventive
measures include restricted access, disinfection of equip –
ment, and quarantine of newly acquired hares. As subclin –
ically infected hares may shed virus, serological screening
of quarantined hares may be advisable. Only seronegative
hares should be permitted entry into the colony . Colonies
with the disease should be quarantined and depopulated.
No vaccine has been developed for use in hares.
Rabbit Calicivirus (RCV)
HISTORY
Non-pathogenic rabbit caliciviruses were first isolated
from rabbits ( Oryctolagus cuniculus ) in a rabbitry in Italy
that seroconverted to RHDV without exhibiting clinical
signs of disease ( Capucci et al., 1996 ). Serological stud –
ies revealed rabbit populations in Europe with serologi –
cal evidence of RHDV , but no clinical evidence of disease
(Capucci et al., 1996, 1997 ; Forrester et al., 1997; Rodak,
1990). In addition, retrospective studies demonstrated
serological evidence of anti-RHDV antibodies prior to
the spread of RHDV virus in Europe ( Moss et al., 2002 ;
Rodak, 1990a), Australia, and New Zealand ( Bruce and
Twigg, 2004; Nagesha et al., 2000; Robinson et al., 2002 ).
Investigators in Italy isolated a non-pathogenic calicivirus,
rabbit calicivirus (RCV), from domestic rabbits ( Capucci
et al., 1996 ). Partial sequences of apparently non-patho –
genic caliciviruses were detected in rabbits ( Oryctolagus
cuniculus ) from Lambay Island, Eire in 2007 ( Forrester
et al., 2007 ). In 2009, another apparently non-pathogenic
calicivirus, RCV-A1, was isolated from wild rabbits
(Oryctolagus cuniculus ) in Australia ( Strive et al., 2009 ).
ETIOLOGY
RCV is considered a strain of RHDV ( Fauquet et al.,
2005 ). Currently , similar viruses have been isolated from

II. RABBITS
14. VIRAL DISEASES
398
Italy (RCV), Eire (Lambay Island strain), and Australia
(RCV-A1) ( Capucci et al., 1996; Forrester et al.,
2007; Strive et al., 2009 ). Genome organization is similar
to RHDV and EBHSV in that there are two open reading
frames as opposed to three which are found in other cali –
civiruses ( Capucci et al., 1996; Strive et al., 2009 ). RCV is
most closely related to RHDV with an amino acid iden –
tity of 91% of the capsid protein, VP60. The amino acid
identity to EBHSV was 75% in the same study ( Capucci
et al., 1996 ). The average identity of the full-length
sequence of RCV-A1 when compared to RHDV was 87%.
Phylogenetic analysis of RCV , the Lambay Island strain,
and the Ashington strain (England) suggests that these
isolates are more closely related to each other than to other
strains of RHDV ( Capucci et al., 1996; Forrester et al., 2007;
Strive et al., 2009 ). Interestingly , the Ashington strain was
isolated from a rabbit ( Oryctolagus cuniculus ) with typical
signs of RHDV . Antigenically , sera from rabbits infected
with RCV cross-reacted with anti-RHDV antibodies, but
not anti-EBHSV antibodies ( Capucci et al., 1996 ). Sera
from ten of 11 healthy rabbits on Lambay Island, Eire, also
cross-reacted with antibody to RHDV ( Forrester et al.,
2007 ) and RT-PCR detected virus sequences shown to be
closely related to RCV from two serum samples.
EPIDEMIOLOGY
RCV-A1 RNA was recovered from the small and large
intestines of wild Eurpean rabbits ( Oryctolagus cunicu –
lus) as well as the fecal pellets from the distal colon,
suggesting a fecal–oral method of transmission similar
to RHDV ( Strive et al., 2009 ). Viral RNA was detected
in Peyer’s patches, spleen, and less frequently in liver.
Experimental inoculation of Oryctolagus cuniculus with
RCV by oronasal route resulted in seroconversion by day
6 after inoculation ( Capucci et al., 1996 ). Uninoculated
cagemates of infected rabbits were seropositive 1–2 days
later, presumably from contact. Rabbits housed in the
same room that had no contact with the infected rabbits
did not seroconvert. RT-PCR detected viral antigens from
the intestine of infected rabbits 3, 5, 6, and 7 days after
inoculation but not from the liver or spleen. Western
blot analyses of intestinal extracts were also positive.
Infection with RCV provided complete protection from
challenge with RHDV . Inoculation of hares with RCV
resulted in no seroconversion and 5 days after inocula –
tion all organs were negative for RCV by RT-PCR. Hares
previously inoculated with RCV and then challenged
with EBHSV were not protected from disease.
CLINICAL SIGNS
Experimental inoculation of Oryctolagus cuniculus with
RCV resulted in no clinical signs of disease ( Capucci et
al., 1996 ) and no clinical signs of disease were observed in
rabbitries with positive serology ( Capucci et al., 1997 ). No
clinical signs were observed in wild rabbits ( Oryctolagus cuniculus ) positive for the Lambay Island strain ( Forrester
et al., 2007 ) or RCV-A1 ( Strive et al., 2009 ).
PATHOLOGY
Gross or histopathologic lesions have not been
described.
DIAGNOSIS
Seroconversion to RHDV without clinical signs of
disease is suggestive of RCV infection ( Capucci et al.,
1997 ). The virus can be detected by RT-PCR of intestinal
contents or feces ( Capucci et al., 1996; Forrester et al.,
2007; Strive et al., 2009 ).
CONTROL
Serologically, RCV is difficult to definitively distin –
guish from RHDV. RCV may represent an older non-
pathogenic lagovirus that provides protection from
RHDV and thus explains why in some areas serore –
activity to RHDV occurs without evidence of disease.
Michigan Rabbit Calicivirus (MRCV)
In January, 2001, an outbreak of hemorrhagic dis –
ease with a 32.5% case fatality rate occurred in a pri –
vate New Zealand White rabbitry in Michigan (Bergin
et al., 2009). Clnical signs consisted of acute fatality,
inappetance, vulvar hemorrhage, conjunctival conges –
tion, opisthotonus, and cyanosis of the lips and ear tips.
On gross necropsy, evidence of icterus, gastric pete –
chiae and ecchymoses, colonic serosal hemorrhage, and
multifocal hemorrhage in the caudal lung lobes was
observed. Histopathology revealed multifocal random
or periportal hepatocellular necrosis and inflammation,
pulmonary and uterine hemorrhages with fibrin clots,
bile duct proliferation, and periductal fibrosis. RT-PCR
of tissue samples ruled out RHDV , however, positive
hepatocytes were detected by immunohistochemistry
utilizing RHDV-specific antibodies.
Further amplification of sequences from pooled
liver samples demonstrated a calicivirus with cap –
sid sequences that most closely aligned to RCV (91.7%
similarity) but were also closely related to RHDV (89.8–
91.3% similarity). When ORF-1 polypepetide genomic
sequence was aligned, excluding the capsid sequences,
MRCV was 77.9–78.5% identical to RHDV , although
other strains of RHDV share similarities of 87.9–98.1%
in these same comparisons. These sequences of RCV
have not been determined (Bergin et al., 2009).
Inoculation of homogenized liver from infected rab –
bits into naïve, SPF (free of Pasteurella sp.) rabbits did
not induce disease after 7 days, although inoculated
rabbits had viral RNA in the liver detected by RT-PCR
and in situ hybridization. This short time course did not
allow for determination of seroconversion (Bergin et al.,
2009).

399 RNA VIRUS INFECTIONS
II. RABBITSClinical signs of disease resolved and rabbits were
depopulated approximately 2 months after the ini –
tial outbreak. Antibody against MRCV was detected in
depopulated rabbits and 20% had gross and non-specific
histopathological changes in their livers consisting of
biliary hyperplasia and periductal to bridging portal to
portal fibrosis. How the virus was introduced into the
colony was not determined (Bergin et al., 2009).
Rabbit Vesivirus
A novel member of the family Caliciviridae was iso –
lated from pooled intestinal contents of five European
rabbits ( Oryctolagus cuniculus ) that died with intestinal
disease in Oregon ( Martín-Alonso et al., 2005 ). Coccidia
were detected in two rabbits and Escherichia coli was
cultured from the pooled intestinal contents. Calicivirus
particles were observed by electron microscopy and a
calicivirus was isolated on porcine kidney cell lines and
partially sequenced. The virus was not neutralized with
anti-calicivirus antibodies (from 40 different antisera)
and sequence analysis indicated it is a novel member
of the Vesivirus genus. The genome consists of a single-
stranded positive sense RNA of 8295 nucleotides and
contains a 2.6-kb subgenomic RNA. Its genomic organi –
zation is unlike lagoviruses, as it contains three ORFs,
rather than two. Phylogenic analysis grouped it in a
clade containing marine caliciviruses and non-human
primate calicivirus, Pan 1.
Paramyxovirus Infections
Rabbit syncytium virus was isolated in chicken
embryos inoculated with extracts of liver and spleen
from a wild Eastern Cottontail rabbit ( Sylvilagus florida –
nus) in Virginia ( Morris et al., 1965 ). The agent causes
a cytopathic effect and syncytia in monkey and ham –
ster kidney cell cultures. Experimentally, suckling mice
are susceptible to the virus, but weaned mice, guinea
pigs, domestic European and Eastern Cottontail rab –
bits failed to develop signs or lesions, although anti –
bodies developed, following inoculation. Sera from
eight of 25 (32%) Eastern Cottontail rabbits trapped in
the same area as the original rabbit had antibodies to
the virus. Antibodies to the virus were not detected in
sera from seven other species trapped in the same area,
Oryctolagus cuniculus , or humans. The virus resembles
the paramyxoviruses in size, nucleic acid type, and in
ether and heat sensitivity. Neither hemagglutination
nor hemadsorption were observed, and the ultrastruc –
ture of the virus has not been described. The virus is
serologically distinct from the known paramyxoviruses.
Evidence of another paramyxovirus, Sendai virus , has
been found in domestic European rabbits ( Oryctolagus
cuniculus ). Sendai virus is the type species of the genus
Respirovirus , family Paramyxoviridae (Fauquet et al., 2005 ) and a primary pathogen of mice ( Percy and Barthold,
2007 ). A serological survey of 23 breeding and labora –
tory colonies, in metropolitan Tokyo and Ibaragi, Chiba,
Saitama, Kanagawa, Shizuoka, and Nagano prefectures,
revealed that 85 of 160 (53%) Japanese White or New
Zealand White rabbits had antibodies to Sendai virus (Iwai
et al., 1986 ). Ito et al. (1987) also detected antibodies to
Sendai virus in rabbit sera in studies comparing antigenic
relationships among paramyxoviruses. Intranasal inocula –
tion of Oryctolagus cuniculus with Sendai virus resulted in
infection as rabbits shed virus for 3–7 days after inocula –
tion, and viral antigen was detected by immunofluores –
cence in the nasal cavities ( Machii et al., 1989 ). Although
rabbits showed no clinical signs and had only a moder –
ate increase of goblet cells in the nasal epithelium, anti –
bodies to the virus developed. One of three uninoculated
rabbits exposed to inoculated rabbits acquired infection.
These studies suggest that Oryctolagus cuniculus is sus –
ceptible to Sendai virus and that this or a related virus may
be endemic in rabbit colonies. Experimentally , genetically
modified Sendai viruses have been used in gene therapy
in rabbits ( Nakamura et al., 1998 ).
Bunyavirus Infections
Evidence of infection with several viruses of the
genus Orthobunyavirus in the family Bunyaviridae has
been detected in leporids. Strains of California encepha –
litis virus found to infect rabbits or hares include
California encephalitis, Snowshoe hare, Tahyna, and
lnkoo viruses ( Fauquet et al., 2005 ). Similar evidence
of infection has been found with viruses of the genus
Bunyamwera virus including cache valley virus, Tensaw
virus, and Northway virus. One other member of the
Bunyaviridae that infects rabbits and is not assigned to a
genus is the Silverwater virus.
Antibodies to California encephalitis virus were ini –
tially detected in Eastern Cottontail rabbits ( Sylvilagus
floridanus ) and Black-tailed jackrabbits ( Lepus califor –
nicus ) in California ( Hammon and Reeves, 1952 ). The
first virus of the group to be recovered from leporids
was the Snowshoe hare virus, isolated in 1959 from
the blood of a sick Snowshoe hare ( Lepus americanus )
in western Montana ( Burgdorfer et al., 1961 ). The virus
is widespread in Snowshoe hare populations of North
America, as serological surveys revealed prevalences
of 40–97% in adult hares ( Hoff et al., 1969; McLean
et al., 1975; Mean, 1983; Newhouse et al., 1963; Yuill et
al., 1969 ). The virus has been isolated from seven spe –
cies of boreal forest mosquitoes, including 0.04% of Aedes
communis (McLean, 1983), and also from the rabbit tick,
Haemaphysalis leporis-palustris (Newhouse et al., 1963 ).
The European hare ( Lepus europeaus ) and wild
European rabbit ( Oryctolagus cuniculus ) appear to be
the major reservoirs of Tahyna virus in Europe ( Bardos,

II. RABBITS
14. VIRAL DISEASES
400
1965, 1975; Danielova et al., 1969; Hannoun et al., 1969;
Simkova, 1963 ). Clinical disease in infected hares has not
been reported. Experimental infection of European rab –
bits results in viremia and antibody formation without
clinical disease ( Hammon and Sather, 1966; Simkova,
1962 ). Thus, domestic European rabbits are useful as sen –
tinels of viral activity since they develop antibodies and
can serve as a source for recovery of virus ( Kolman et al.,
1966; McKiel et al., 1966 ). A serological survey in Finland
revealed that 5% of Mountain hares ( Lepus timidus ) and
none of the Lepus europeaus tested had antibodies against
Inkoo virus ( Brummer-Korvenkontio, 1973 ). Similar
viruses have been isolated from mosquitoes and humans
in Russia ( Vanlandingham et al., 2002 ).
Cache valley virus, Tensaw virus, and Northway
virus are mosquito-borne viruses of North America.
Cache valley virus has a wide geographic distribu –
tion in North America ( Blackmore and Grimstad, 2008 ),
whereas Tensaw virus is found predominantly in the
Southeastern United States ( Bigler et al., 1975 ) and
Northway virus in Alaska and California ( Campbell
et al., 1990, 1991; Walters et al., 1999; Zarnke et al., 1983 ).
Recent evidence suggests these host ranges may not be
as restrictive as earlier thought ( Sahu et al., 2002 ). The
amplifying hosts for all of these viruses are unclear, how –
ever antibodies recognizing Cache valley virus have
been found in Desert cottontails ( Sylvilagus audubonii ),
Eastern cottontails ( Sylvilagus floridanus ), and Black-tailed
jackrabbits ( Lepus californicus ) (Blackmore and Grimstad,
2008 ). Antibodies to Tensaw virus have been found in
the Swamp rabbit ( Sylvilagus acquaticus ) (Calisher et al.,
1986 ), Marsh rabbit ( Sylvilagus palustris ) (Bigler et al.,
1975 ), and New England cottontail rabbit ( Sylvilagus tran –
sitionalis ) (Sudia et al., 1969 ) and those to Northway virus
in the Snowshoe hare ( Lepus americanus ) (Zarnke et al.,
1983 ). Experimental infection of Eastern cottontail rabbits
with Cache valley virus resulted in transient viremia of
insufficient magnitude to infect mosquitoes ( Coquillettidia
perturbans ) suggesting that this species may not be an
important vertebrate host for this virus ( Blackmore and
Grimstad, 2008 ). In contrast, rabbits and hares may be
important hosts for Tensaw and Northway viruses as
experimental infection of New England cottontail rab –
bits ( Sylvilagus transitionalis ) with Tensaw virus resulted
in viremia of sufficient magnitude to infect mosquitoes
(Anopheles quadrimaculatus ) (Sudia et al., 1969 ). Living
near Snowshoe hares ( Lepus americanus ) was found to be
a significant risk factor for human serologic reactivity to
Northway virus ( Walters et al., 1999 ).
Silverwater virus has been isolated from Snowshoe hares
(Lepus americanus ) and from rabbit ticks ( H. leporis palus –
tris) from Snowshoe hares in Ontario and Alberta, Canada
(McLean and Larke, 1963; Yuill et al., 1969 ). Although no
disease has been reported in infected hares, they appear to
play a central role in the natural cycle of the virus.Togavirus Infections
Serological evidence of infection with Western (WEE),
Eastern (EEE), and Venezuelan (VEE) equine encephalitis
viruses , members of the genus Alphavirus , in the family
Togaviridae (Fauquet et al., 2005 ) has been found in rab –
bits and hares. Wild Sylvilagus as well as hares ( Lepus
californicus and Lepus americanus ) have antibodies to
WEE ( Bowers et al., 1969; Yuill et al., 1969 ). Infection
studies of Oryctolagus cuniculus with strains of WEE
isolated from South America and North America dem –
onstrated virus strain variation in clinical signs and
mortality ( Bianchi et al., 1997 ). A South American strain
(AG80-646) resulted in early onset of anorexia and rear
limb paralysis with rabbits dying 6 days after inocula –
tion. The North American strain caused no detectable
clinical signs, but the rabbits seroconverted. Antibodies
to EEE ( Yuill et al., 1969 ) and VEE ( Hoff et al., 1970 )
have been detected in Snowshoe hares ( Lepus ameri –
canus ) and antibodies to VEE have been detected in
Eastern cottontail rabbits ( Sylvilagus floridanus ) (Smart
and Trainer, 1975 ).
Complement fixation, hemagglutination-inhibition,
ELISA, and plaque reduction neutralization tests are
commonly used for diagnosis of these infections in
humans and animals, although cross-reactivity between
different viruses is problematic with all of these tests
(O. I. E., 2008). An epitope-blocking assay was devel –
oped and tested on rabbit sera to better separate the
individual virus groups (Pässler and Pfeffer, 2003). The
infectious agent can be identified by viral isolation or
PCR (O. I. E., 2008; Vodkin et al., 1993).
Flavivirus Infections
Two members of the Japanese encephalitis group
in the family Flaviviridae , genus Flavivirus have been
shown to infect rabbits or hares, St. Louis encephalitis
virus and West Nile virus (WNV) and one member of
the mammalian tick-borne virus group of the genus
Flavivirus has been shown to infect hares, Powassan virus
(Fauquet et al., 2005 ). Antibodies to St. Louis encepha –
litis virus have been detected in Snowshoe hares ( Lepus
americanus ) (Yuill et al., 1969 ). Experimentally, Eastern
cottontail rabbits ( Sylvilagus floridanus ) have been
shown to be susceptible to WNV infection ( Tiawsirisup
et al., 2005 ). WNV is a mosquito-transmitted pathogen
of significant public health importance ( Hayes et al.,
2005 ). The major route of infection of humans is by the
bite of a mosquito infected by ingesting a blood meal
from an infected bird. In general, mammalian hosts of
WNV are dead-end hosts and do not develop viremia
of sufficient magnitude to infect mosquitoes ( Bowen
and Nemeth, 2007 ). However, certain species, including
Eastern cottontail rabbits ( Sylvilagus floridanus ), develop
viremia which infects mosquitoes ( Culex pipiens and

401
II. RABBITSREFERENCES
Culex salinarius ) and therefore, may contribute to the
endemic cycle of WNV in North America ( Tiawsirisup
et al., 2005 ). Infection in rabbits occurs without clini –
cal signs. Powassan virus is a tick-borne virus of emerg –
ing public health concern in the United States, Canada,
and Russia ( Ebel, 2010 ). Antibodies to Powassan virus
have been found in Snowshoe hares ( Lepus americanus )
(McLean et al., 1961; Zarnke and Yuill, 1981 ). Snowshoe
hares experimentally infected with the virus develop
no clinical symptoms, but become viremic for 1–3 days.
Other vertebrates, including red squirrels ( Tamiasciurus
hudsonicus ), chipmunks ( Tamias amoenus ), groundhogs
(Marmota monax ), and white-footed mice ( Peromyscus
leucopus ) may be more important in perpetuating the
virus than Snowshoe hares ( Ebel, 2010 ).
Picobirnavirus Infections
A rabbit Picobirnavirus , a member of the Picobirnaviridae
family , was detected in rabbit feces in a study in which
rabbits were inoculated with human Picobirnavirus
(Fregolente et al., 2009; Gallimore et al., 1993 ).
Picobirnaviruses are non-enveloped bisegmented, double-
stranded RNA viruses of 35–40 nm in diameter. Members
of this family of viruses have been found in the feces of
many different hosts, including humans, pigs, rabbits,
dogs, rats, snakes, and birds ( Fregolente et al., 2009 ). They
are thought to be opportunistic gastrointestinal patho –
gens associated with clinical disease in humans ( Giordano
et al., 1999 ). These viruses have been associated with gas –
trointestinal disease in HIV-infected humans ( Giordano
et al., 1999 ; Grohmann et al., 1993) and may be important
pathogens or co-pathogens in immunosuppressed rab –
bits. The picobirnavirus genome detected in rabbits con –
sisted of two segments; 2.3 kbp and 1.85 kbp ( Gallimore
et al., 1993 ). The first segment contains one major ORF
and two smaller ORFs and has been partially sequenced
(Green et al., 1999 ). This segment is thought to encode
a capsid protein while the second segment appears to
encode an RNA-dependent RNA polymerase ( Rosen
et al., 2000 ). Picobirnaviruses do not replicate in tissue
culture, although they were successfully detected in the
feces of rabbits inoculated orally with rabbit Picobirnavirus
(Gallimore et al., 1993; Ludert et al., 1995 ). No clinical
signs of disease resulted, however, anti- Picobirnavirus anti –
bodies developed ( Gallimore et al., 1993 ).
Rabies Virus Infections
Rabies virus, the type species of the genus Lyssavirus
and family Rhabdoviridae (Fauquet et al., 2005 ), is a neu –
rotropic virus that infects mammals resulting in a gen –
erally fatal, progressive encephalomyelitis ( Manning
et al., 2008 ). It is found in high titers in the saliva of clinically ill animals and is typically transmitted
through a bite. Rabies virus infections in rabbits and
hares are rare, as these species often do not survive
encounters with rabid animals. From 1971 to 1984,
seven rabid rabbits were reported to the CDC, includ –
ing four Eastern cottontails ( Sylvilagus floridanus )
(Fishbein et al., 1986), while in the next 9 years, 17 cases
of rabies in rabbits were reported to the CDC ( Childs
et al., 1997 ). Of these 17, 11 were domestic European
rabbits ( Oryctolagus cunniculus ) and the others were
unknown. Four viruses were typed and shown to be
either raccoon or skunk variants of rabies virus. In 1995,
raccoon-variant rabies was reported in seven domes –
tic European rabbits in New York State ( Eidson et al.,
2005 ). Rabies is typically diagnosed by fluorescent anti –
body test on brain tissue, however, strains may be fur –
ther identified using monoclonal antibodies or genetic
means such as nucleic acid probes, PCR, and DNA
sequencing (O. I. E., 2008).
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