KVÄLLSSYMPOSIUM 2008 Vaccinering av hund och katt

KVÄLLSSYMPOSIUM 2008
Vaccinering av hund och katt
Stockholm 25 november – Göteborg 26 november – Lund 27 november
Kennelhosta – vad är det?
Jessica Ingman, Bitr. Statsvet., Sekt. f.
häst, hund och katt, SVA
Nobivac KC® vet. – nytt vaccin från
Intervet/Schering-Plough Animal
Health
Agneta Gustafsson, Teknisk chef,
I/SPAH
Svenska erfarenheter av
Nobivac KC® vet.
Anette Johansson, Distriktsvet., Kiruna
Presentation av SVAs rabiestiterstudie
Louise Treiberg-Berndtsson, Bitr. Statsvet.,
Sekt. f. virologisk diagnostik, SVA
Kort bensträckare
Nobivac® Tricat Novum vet. – nytt
vaccin från Intervet/Schering-Plough
Animal Health
Anna-Karin Lieber, Produktchef, I/SPAH
Vaccination av hund och katt 2008
Ulrika Windahl, Bitr. Statsvet., Sekt. f.
häst, hund och katt, SVA
Paneldiskussion med föredragshållarna
Intervet AB
Box 6103
102 33 Stockholm
Tel 08–522 216 60
www.intervet.se

Kennelhosta – vad är det?
Bitr. statsveterinär Jessica Ingman
Enheten för djurhälsa och antibiotikafrågor, Statens veterinärmedicinska anstalt
Kennelhosta
• Är ett samlingsbegrepp för kikhosteliknande symtom hos hund.
• Akut, mycket smittsam luftvägsinfektion som karakteriseras av akut insättande
hostattacker (kväljningar/slem/nosflöde).
• Multifaktoriell etiologi.
• Graden av symtom kan variera beroende på vilket eller vilka agens som är
inblandade.
• Infektion med enbart ett agens ger oftast lindrigare symtom.
• Prognosen god vid okomplicerade infektioner, men immuniteten kan vara
kortvarig.
Smittspridning
• Infektion sprids snabbt och effektivt bland hundar som hålls tillsammans i
större grupper.
• Smitta direkt nos-nos-kontakt, hosta/nysningar, indirekt via händer/föremål.
• Kliniska symtom uppträder vanligtvis inom 3-5 dagar efter infektion, även om
inkubationstid för kennelhosta anges till 3-10 dagar.
• För de vanligaste virala agens kan utsöndring av virus ske upptill två veckor
efter infektion.
• För B. bronchiseptica kan dock utsöndring pågå betydligt längre tid.
Etiologi
• Virus
- Hundens parainfluensavirus typ 2 (CPiV-2)
- Hundens adenovirus typ 2 (CAV-2)
- Hundens herpesvirus typ 1 (CHV-1)
- Canine respiratory corona virus (CRCoV)
- (valpsjuka…)
• Bakteriella agens
- Bordetella bronchiseptica
- Mykoplasma-arter, streptokocker, pasteurella-arter, koliforma bakterier och
pseudomonas-bakterier har också påvisats hos hostande hundar, men då
främst som sekundärinfektion eller eventuellt bifynd.
• Hundens parainfluensavirus typ 2 och Bordetella bronchiseptica är de agens som
enligt litteraturen oftast isoleras från hundar med infektiös trakeobronkit, men
flera andra virus och bakterier kan påverka symtombild och sjukdomsförlopp.

Hundens parainfluensavirus typ 2
• CPiV-2, canine parainfluenza virus
• RNA-virus, familjen Paramyxoviridae
• Det vanligaste virus som isoleras från luftvägarna hos hund i samband med
kennelhostesymtom.
• Orsakar vid ensam infektion vanligen en relativt lindrig infektion i övre
luftvägarna med kortvarig, övergående hosta och minimal allmänpåverkan.
• (Katter kan bli subkliniskt infekterade och utsöndra virus – okänt dock om
betydelse för smittspridning hos hund.)
Hundens adenovirus typ 2
• Virus som är nära släkt med det virus (CAV-1) som orsakar HCC, men CAV-2
angriper huvudsakligen luftvägarna.
• Infektion orsakar vanligen inga påtagliga kliniska symtom.
• De flesta hundar är idag vaccinerade mot CAV-2 (ger skydd mot CAV-1 som
orsakar HCC).
Hundens herpesvirus typ 1
• Är ett virus som också angriper de övre luftvägarna hos hund, men som
vanligtvis inte orsakar några tydliga kliniska symtom. Anses inte längre vara
någon väsentlig orsak till kennelhosta.
• (Har betydelse vid dödlig infektion hos nyfödda valpar.)
CRCoV
• Canine respiratory corona virus
• Relativt nyupptäckt virus som isolerats från hundar med kennelhostesymtom.
• Är olikt från CCoV – den enteriska formen
• Har oftast associerats med lindriga kliniska symtom.
Valpsjukevirus
• CDV, canine distemper virus
• Kan ge respiratoriska symtom och har därför tidigare beskrivits i samband med
kennelhostekomplexet, men har idag mindre betydelse pga. utbredd vaccination.
B. bronchiseptica
• Gramnegativ, aerob bakterie.
• Kan förekomma i luftvägarna även hos friska individer.
• Hundratals olika isolat påvisade med olika virulens och patogenicitet.
• I experimentella studier orsakar infektion med enbart B. bronchiseptica typiska
symtom på kennelhosta framför allt hos unga hundar.
• Bakterien kan utsöndras från övre luftvägarna i 3-4 månader efter
genomgången infektion.

Sammanfattningsvis om kennelhosta
• Kennelhosta är ett komplext komplex - flera möjliga agens, multifaktoriell
etiologi.
• Patogena agens kan vara olika vid olika utbrott.
• En hund som haft kennelhosta är inte skyddad mot att få sjukdomen igen vid
ett senare tillfälle.

Vet. Res. 38 (2007) 355–373 355
c○ INRA, EDP Sciences, 2007
DOI: 10.1051/vetres:2006058
Review article
Canine respiratory viruses
Canio Buonavoglia*, Vito Martella
Department of Animal Health and Wellbeing, Faculty of Veterinary Medicine of Bari, Italy
(Received 27 April 2006; accepted 28 August 2006)
Abstract – Acute contagious respiratory disease (kennel cough) is commonly described in dogs
worldwide. The disease appears to be multifactorial and a number of viral and bacterial pathogens
have been reported as potential aetiological agents, including canine parainfluenza virus, canine
adenovirus and Bordetella bronchiseptica, as well as mycoplasmas, Streptococcus equi subsp.
zooepidemicus, canine herpesvirus and reovirus-1,-2 and -3. Enhancement of pathogenicity by multiple
infections can result in more severe clinical forms. In addition, acute respiratory diseases
associated with infection by influenza A virus, and group I and II coronaviruses, have been described
recently in dogs. Host species shifts and tropism changes are likely responsible for the
onset of these new pathogens. The importance of the viral agents in the kennel cough complex is
discussed.
kennel cough / respiratory disease / dogs / viruses
1.
Table of contents
Introduction ......................................................................................................355
2. Canineadenovirustype2 .....................................................................................356
3. Canineherpesvirus .............................................................................................357
4. Canineinfluenzavirus ..........................................................................................359
5. Canineparainfluenzavirus ....................................................................................360
6. Caninereovirus..................................................................................................362 7. Caninerespiratorycoronavirus ..............................................................................363
8. Pantropiccaninecoronavirus.................................................................................364 9. Conclusions ......................................................................................................365
1. INTRODUCTION
Infectious tracheobronchitis (ITB) or
kennel cough is the term used by veterinarians
to describe an acute, highly contagious
respiratory disease in dogs affecting the
larynx, trachea, bronchi, and occasionally
the nasal mucosa and the lower respiratory
tract [6].
* Corresponding author:
c.buonavoglia@veterinaria.uniba.it
Mild to severe episodes of cough and
respiratory distress are characteristic clinical
features recognized in affected dogs.
ITB has worldwide distribution and is recognized
as one of the most prevalent infectious
diseases of dogs. The disease is frequently
described in dogs housed in groups
in rehoming centers and boarding or training
kennels.
Two clinical forms of ITB have been described.
The uncomplicated form is most
Article available at http://www.edpsciences.org/vetres or http://dx.doi.org/10.1051/vetres:2006058

356 C. Buonavoglia, V. Martella
common and is characterized as a dry,
hacking cough, often in association with
gagging and retching behavior. The dogs
are affected by a self-limiting, primarily
viral infection of the trachea and bronchi.
A complicated form of ITB is described
in puppies or immuno-compromised dogs.
In the complicated forms, secondary bacterial
infections and involvement of pulmonary
tissue overlaps the viral process.
The cough is associated with mucoid discharges.
The condition may progress to
bronchopneumonia and, in the most severe
instances, to death [6].
Multiple agents, bacterial and viral, are
implicated in the aetiology of ITB. Coisolation
of viral and bacterial pathogens
is frequent, while experimental infections
with single pathogens may result in subclinical
or mild forms of disease, suggesting
a multi-factorial pathogenesis. Many
agents likely play a role in ITB, such
as canine parainfluenza virus [2], canine
adenovirus [55], Bordetella bronchiseptica
[18], and mycoplasmas [36, 128].
Streptococcus equi subsp. zooepidemicus,
has been associated with severe to fatal
respiratory forms in dogs alone or in
mixed infections [36, 61, 173]. Recently,
outbreaks of influenza A virus, initially
misdiagnosed as ITB, have been reported
in the USA [45,173]. In addition, novel canine
coronaviruses, a pantropic variant of
CCoV type II [29] and the canine respiratory
coronavirus virus, CRCoV [60], have
been detected from the respiratory tract
of either symptomatic or asymptomatic
dogs. Canine herpesvirus and reovirus-1,-
2,-3 have rarely been reported from dogs
with kennel cough but are not thought
to play a major role in the disease complex
[86,97]. Vaccines are available against
some of these infectious agents but regular
vaccination in kennels often fails to prevent
ITB.
An overview of the viral agents that
have been associated with ITB in dogs,
with particular regards to newly described
viruses, is reported.
2. CANINE ADENOVIRUS TYPE 2
Canine adenovirus type 2 (CAV-2) determines
unapparent to mild infection of
the respiratory tract and is regarded as one
of the causes of the common widespread
ITB [154]. CAV-2 has also been implicated
in episodes of enteritis [76, 98] and has
been detected in the brain of dogs with
neurological signs [19].
The virus was first detected in 1961,
in Canada, from dogs affected by laryngotracheitis
[55]. The isolate, strain Toronto
A26/61, was characterized as an adenovirus,
and was initially considered to be
an attenuated strain of canine adenovirus
type 1 (CAV-1). Subsequently, structural
and antigenic differences were observed
and strain A26/61 was proposed as the
prototype of a distinct canine adenovirus,
designated as type 2 (CAV-2) [65, 75, 104,
107, 152, 172]. CAV-1 and CAV-2 were
found to be genetically different by restriction
endonuclease analysis [10, 75] and by
DNA hybridization [106]. The complete
sequence analysis of both the CAV-1 and
CAV-2 genome has revealed about 75%
nucleotide identity [49, 114]. Although
CAV-1 and CAV-2 are related genetically
and antigenically [109,168], they have different
tissue tropism. Vascular endothelial
cells and hepatic and renal parenchymal
cells are the main targets of CAV-1, while
the respiratory tract epithelium and, to a
limited degree, the intestinal epithelium,
are the targets of CAV-2 [3, 140, 153]. In
addition, the two types display different
hemagglutination patterns [105].
Infection with CAV-2 appears to be
widespread in dogs that are not immune to
CAV-1 or CAV-2. CAV-2 was isolated from
34 out of 221 throat swabs of pups with and
without respiratory signs that were taken
to a veterinarian for vaccination [4]. Likewise,
pups in pet shops and in laboratory

animal colonies were found to carry CAV-2
in the respiratory tract [6,20]. By converse,
CAV-2 was not detectable in dogs vaccinated
in a rehoming center [61].
The host range of CAV-2 includes
a broad number of mammalian species.
Wild-life animals may be a source of infection
for domestic dogs. The overall
prevalence of antibodies to canine adenoviruses
in European red foxes (Vulpes
vulpes) in Australia was 23.2% with
marked geographical, seasonal and age differences
[134], while the prevalence of antibodies
was 97% in Island foxes (Urocyon
littoralis) in the Channel Islands, California
[70]. Antibodies to CAV-2 were also
detected in free-ranging terrestrial carnivores
and in marine mammals in Alaska
and Canada, including black bears (Ursus
americanus), fishers (Martes pennanti),
polar bears (Ursus maritimus), wolves
(Canis lupus), walruses (Odobenus rosmarus)
and Steller sea lions (Eumetopias
jubatus) [30, 120, 147].
The route of infection of CAV-2 is oronasal.
The virus replicates in non-ciliated
bronchiolar epithelial cells, in surface cells
of the nasal mucosa, pharynx, tonsillar
crypts, mucous cells in the trachea and
bronchi in peribronchial glands and type
2 alveolar epithelial cells. In addition to
these tissues, the virus can be isolated
from retropharyngeal and bronchial lymph
nodes as well as from the stomach and
the intestine. The peak of replication is
reached by 3–6 days post infection. Subsequently,
virus loads rapidly decline, in
relation to the production of antibodies,
and CAV-2 usually can not be isolated by
9 days post infection. Respiratory signs
are consistent with damage of bronchial
epithelial cells. There may be evidence
of narcotising bronchitis or bronchiolitis
and of bronchiolitis obliterans. Infection of
type 2 alveolar cells is associated with interstitial
pneumonia [5, 6, 9, 35, 47, 90].
Dogs exposed to CAV-2 alone rarely
show spontaneous disease signs, although
Canine respiratory viruses 357
lung lesions can be extensive. When additional
bacterial or viral agents are involved,
the ITB complex can be observed [5, 6].
Antibodies to CAV-2 antigens have
been demonstrated by hemagglutinationinhibition,
agar gel diffusion, virus precipitation,
complement fixation and by neutralization
[90]. Protection appears to correlate
with the neutralizing antibody levels [3,8].
Nasal or throat swabs appear to be suitable
for virus isolation. Primary dog kidney
cells have been used successfully for
isolation and cultivation of CAV-2 [55].
However, a variety of cell lines are similarly
susceptible to CAV-2 and to CAV-1
[171]. Demonstration of CAV-2 antigen by
immunofluorescence in acetone-fixed lung
sections or tissue imprints is used for diagnosis
of CAV-2. A polymerase chain
reaction (PCR) assay has been developed
to detect canine adenoviruses and to distinguish
between CAV-1 and CAV-2 [82].
Modified live CAV-2 vaccines proved
to be highly effective in reducing the circulation
of CAV-2 in canine populations.
Dogs vaccinated with CAV-2 develop immunity
to both CAV-1 and CAV-2 [3, 8].
In a similar fashion, dogs vaccinated with
CAV-1 develop immunity to both CAV-1
and CAV-2 [43]. However, the use of
CAV-2 for immunization of pups against
both canine adenovirus types has eliminated
safety side-effects encountered with
CAV-1 vaccines, i.e. the occurrence of ocular
lesions [24,48,90]. Maternally-derived
antibodies in pups may prevent active immunization
after vaccine administration up
to the age of 12-16 weeks [8]. Vaccine
administration by the intranasal route has
been proposed to overcome the interference
of maternal antibodies [8], but products
for intranasal vaccination are not marketed.
3. CANINE HERPESVIRUS
Canine herpesvirus (CHV) is a memberoftheAlphaherpesvirinae
subfamily

358 C. Buonavoglia, V. Martella
of the Herpesviridae [159]. CHV was first
described in the mid 1960s from a fatal
septicemic disease of puppies [32]. Infection
of susceptible puppies of less than
two weeks of age may result in fatal generalized
necrotizing and hemorrhagic disease,
while pups older than two weeks and
adult dogs often do not show any clinical
signs [32]. Infection in older dogs appears
to be restricted to the upper respiratory
tract [7]. CHV is also transmitted transplacentally,
resulting in fetal death [77].
Serological surveys have shown a relatively
high prevalence of CHV in household
and colony-bred dogs. The prevalence
of antibodies in dogs was 88% in England,
45.8% in Belgium, and 39.3% in
the Netherlands [129, 133, 135]. Serological
studies in Italy have revealed a similar
prevalence in kennelled dogs (27.9%),
while the prevalence was lower in pets
(3.1%) [142].
The host range of CHV is restricted to
dogs [91]. However, antibodies to CHV
have been detected in the sera of European
red foxes (Vulpes vulpes) in Australia [134]
and Germany [156] and in sera of North
American river otters (Lontra canadensis)
from New York State [88], while a CHVlike
virus has been isolated from captive
coyote pups [64].
CHV appears to be a monotypic virus,
as defined by antigenic comparison of various
isolates [32, 122]. The gene structure
of CHV has yet to be determined, since
no CHV strain has been completely sequenced
and only a few genes have been
identified [73, 96, 132]. Restriction mapping,
southern blot hybridization and sequence
analysis have shown that the overall
structure of CHV resembles those of
other alphaherpesviruses and that CHV is
genetically related to feline herpesvirus
(FHV-1), phocid herpesvirus 1 and to the
equid herpesviruses 1 and 4 [103,130,138,
169].
Like other herpesviruses, attachment of
CHV to permissive cells (MDCK) ap-
pears to be mediated by heparan sulfate,
as observed for FHV and for other herpesviruses
[99, 116].
After both symptomatic and asymptomatic
infections, dogs remain latently infected
and virus may be excreted at unpredictable
intervals over periods of several
months, or years. Reactivation of latent
virus may be provoked by stress (movement
to new quarters, introduction of new
dogs) or, experimentally, by immunosuppressive
drugs (corticosteroids) or antilymphocyte
serum. Latent virus, demonstrated
by the polymerase chain reaction, persists
in the trigeminal ganglia, but other sites
such as the lumbo-sacral ganglia, tonsils,
and parotid salivary gland have been identified
[31, 33, 112, 119].
Canine herpesvirus has been detected in
dogs with ITB [22] but its role remains
controversial. Experimental infection has
been shown to cause mild clinical symptoms
of rhinitis and pharyngitis [7] or to
result in ITB-related disease [85]. Experimental
infection by the intravenous route
in adult foxes results in fever, lethargy and
respiratory signs, while peroral infection
does not [131].
A long-term survey in a population
of dogs in a rehoming center has evidenced
CHV in 9.6% of lung and 12.8%
of tracheal samples. CHV infections occurred
later than other viral infections. In
contrast to CRCoV and CPIV, that were detected
more frequently within the first and
second week, respectively, CHV was detected
more frequently at weeks 3 and 4
after dog introduction in the kennel. Interestingly,
CHV infection was apparently related
to more-severe respiratory signs [61].
Whether the presence of CHV is responsible
for increased disease severity or viceversa
is not clear. In a 1-year study in
training centers for working dogs, seroconversion
to CHV appeared to be more
frequent in dogs infected by CRCoV [62],
a finding that is more consistent with virus
reactivation after disease-induced stress.

An inactivated, subunit vaccine has
been available in Europe since 2003. The
vaccine is specifically indicated for bitches
during pregnancy. The vaccine was shown
to provide good immunity to newborn pups
after two injections had been administered
to their dams.
4. CANINE INFLUENZAVIRUS
Influenza is globally the most economically
important respiratory disease in
humans, pigs, horses, and in the avian
species. Influenza A viruses have enveloped
virions of 80 to 120 nm in diameter,
with about 500 spikes of 10 to
14 nm in length radiating outward from the
lipid envelope [170]. The genome is composed
of eight segments of single-stranded
RNA that segregate independently. The
spike proteins, HA (hemagglutinin) and
NA (neuraminidase), elicit neutralizing antibody
response and provide the basis for a
dual classification system by H (1 to 16)
and N (1 to 9) subtypes [67, 170].
Distribution of the various subtypes is
species-restricted but interspecies transmissions
may occur, notably between
avians and mammalians [170]. H3N2,
H2N2 and H1N1 strains have been responsible
for influenza-associated disease
and mortality in the last decades in humans,
while fatal infections by the highly
pathogenic H7N7 and H5N1 avian strains
have occurred sporadically [41, 66, 151].
In pigs, influenza A infection is caused by
H1N1 and H3N2 subtypes [27]. Two different
subtypes of equine influenza virus,
H7N7 and H3N8, have been associated
with disease in horses [149, 162, 163]. Virtually
all the H and N subtypes have been
signaled in the avian species that act as a
vast, continual reservoir for mammalians
[164, 170].
Until recently, dogs were regarded
unanimously as non-susceptible hosts to
influenza virus A. In the 1980s, antibodies
to human influenza A viruses were
Canine respiratory viruses 359
detected in the sera of dogs by hemagglutination
inhibition and serum neutralization
assays, suggesting a possible exposure of
dogs to human influenza viruses [28].
The first documented evidence of influenza
A in dogs was obtained in 2004 in
the USA, where outbreaks of severe respiratory
disease were reported in Florida
racing greyhounds [45]. Additional outbreaks
of respiratory disease were reported
in 2004 in 6 states and 2005 in 11 states
throughout the USA and the infection was
also confirmed in pet dogs. These cases
occurred in animal shelters, humane societies,
rescue groups, pet stores, boarding
kennels, and veterinary clinics [45, 173].
The viruses were found to agglutinate
chicken erythrocytes and two
strains were isolated in Madin-Darby
canine kidney cells (MDCK) from
lung and bronchioalveolar lavage
fluid [45, 173], A/ca/Florida/43/2004
and A/ca/Iowa/13628/2005. Retrospective
serological investigation demonstrated
that the virus was present before 2004,
but not before 1998, and a strain,
A/eq/Florida/242/03, was isolated from
archival tissues of a greyhound that had
died from hemorrhagic bronchopneumonia
in 2003 [45].
Molecular and antigenic analyses of
influenza viruses isolated from the various
influenza outbreaks in racing greyhounds
revealed that the canine strains are
closely related to H3N8 equine influenza
viruses [45, 173]. The HA and NA genes
of the canine isolates are genetically close
(96%–98% nucleotide identity) to the HA
and NA genes of recent H3N8 equine
influenza viruses. Sequence and phylogenetic
analysis of all the 8 genome segments
indicated that the canine influenza viruses
form a monophyletic group, a finding that
is consistent with a single interspecies
virus transfer, and that the virus likely got
adapted to the canine host by accumulation
of point mutations rather than by exchange
of genome segments via reassortment with

360 C. Buonavoglia, V. Martella
other influenza virus A strains, since all the
genome segments are of equine origin [45].
Two distinct clinical forms have been
described in dogs infected with influenza
virus with illness rates being nearly 100%.
A milder illness is described in most dogs,
which is characterized by initial fever and
then cough for 10 to 14 days, followed
by recovery. The cough is usually moist,
but in some dogs can be dry and resemble
the ITB complex. A thick nasal discharge
may be described, which is usually caused
by a secondary bacterial infection. A peracute
death associated with haemorrhage
in the respiratory tract has been observed
in about 5% of the dogs. The severe form
is accompanied by rapid respiration and
high fever (40–41 ◦ C). Post-mortem examination
of dogs dead after the peracute
form revealed extensive haemorrhage in
the lungs, mediastinum and pleural cavity.
The lungs exhibited extensive red to
red-black discoloration with moderate to
marked palpable firmness. Mild fibrinous
pleuritis was also noted. Histological examination
revealed tracheitis, bronchitis,
bronchiolitis and suppurative bronchopneumonia.
Lung sections were characterized
by severe hemorrhagic interstitial or
bronchointerstitial pneumonia. Patchy interstitial
change with alveolar septal thickening,
coagula of debris in the alveoli, and
associated atelectasis were evident, along
with foci of pyogranulomatous bronchointerstitial
pneumonia and dilatation of airways
by degenerate cells and debris. Scattered
vasculitis and vascular thrombi were
also observed [45, 173]. The disease has
been reproduced by experimental inoculation
of the virus [45].
Therapeutic administration of broadspectrum
antimicrobial drugs reduces the
severity but can not control the disease
[173]. In the milder forms, a thick
green nasal discharge, which most likely
represents a secondary bacterial infection,
usually resolves quickly after treatment
with a broad-spectrum bactericidal antimi-
crobial. In the more severe forms, pneumonia
is likely caused by bacterial superinfection,
and responds best to hydration
and broad-spectrum bactericidal antimicrobials.
No vaccine is available to protect
dogs against canine influenza. Vaccination
against other pathogens causing respiratory
disease, however, may help prevent
more common respiratory pathogens
from becoming secondary infections in
a respiratory tract already compromised
by influenza infection. The canine influenza
virus appears to be easily inactivated
by common disinfectants (e.g., quaternary
ammonium compounds and bleach
solutions). Protocols should be established
for thoroughly cleaning and disinfecting
cages, bowls, and other surfaces between
use, as well as for disinfections of personnel
before and after handling of animals.
There is no rapid test for direct diagnosis
of acute canine influenza virus infection.
Serological assays may detect antibodies
to canine influenza virus as early
as 7 days after onset of clinical signs. In
equipped laboratories, viral isolation on
tissue cultures and reverse transcription
(RT)-PCR or real time PCR analysis may
be applied to fresh lung and tracheal tissues
of dogs that have died from pneumonia and
to respiratory secretion specimens from ill
animals.
5. CANINE PARAINFLUENZAVIRUS
Canine parainfluenzavirus (CPIV) was
first reported in the late 1960s from laboratory
dogs with respiratory disease [2] and
from a sentry dog with respiratory disease
of the upper tract [44]. Subsequent studies
revealed that the virus was frequent in dogs
with respiratory disease [11, 22, 42, 110,
137, 158], suggesting a key role, along
with Bordetella bronchiseptica, in the aetiology
of ITB.
Parainfluenza viruses include important
pathogens of the respiratory tract of mammals
and birds. The term parainfluenza

was originally adopted after the influenzalike
symptoms observed in infected patients
and after the influenza-like hemagglutination
and neuraminidase activities
exhibited by the virus particles. Parainfluenza
viruses are classified in the family
Paramyxoviridae, subfamily Paramyxovirinae.
CPIV is antigenically similar to
the simian virus 5 (SV5) and to porcine,
bovine, ovine and feline parainfluenza
viruses [1, 127]. Sequence analysis of the
fusion protein-encoding gene has revealed
that CPIV has 99.3% nucleotide similarity
to porcine parainfluenza virus, 98.5%
to SV5 and 59.5% nt to human parainfluenza
virus 2 [111]. Accordingly, CPIV
is regarded as a host variant of SV5, within
the genus Rubulavirus and has been tentatively
proposed as parainfluenza virus 5
(PI-5) [39]. Viruses genetically similar to
SV5 have been detected in humans in more
occasions although the relationship to any
human disease remains contentious [39].
The virus is composed of a single
stranded RNA genome of negative polarity
and is surrounded by a lipid envelope
of host cell origin. The genome
of SV5 contains seven genes that encode
eight proteins: the nucleoprotein
(NP), V/phosphoprotein (V/P), matrix
(M), fusion (F), small hydrophobic (SH),
hemagglutinin–neuraminidase (HN), and
large (L) genes [92]. The HN protein
is involved in cell attachment to initiate
virus infection and mediates hemagglutination
[100]. In addition, HN has neuraminidase
activity. The F protein mediates
fusion of the viral envelope with the cell
membrane [143]. The V protein blocks interferon
(IFN) signaling and inhibits IFN
synthesis. Interaction of the virus with the
IFN system is regarded as a critical factor
in the outcome of the infection [23,54,121,
146].
Parainfluenza is highly contagious and
the prevalence of infection appears to be
related to the density of the dog population.
CPIV is excreted from the respiratory tract
Canine respiratory viruses 361
of infected animals for 8-10 days after infection
and is usually transmitted by direct
contact with infected aerosol [6]. The virus
may spread rapidly in kennels or shelters
where a large number of dogs are kept together.
The virus was detected in 19.4%
of tracheal and 9.6% of lung samples of
dogs in a rehoming centre where ITB was
endemic and persisted, in spite of regular
vaccination against canine adenovirus
type-2, distemper and parainfluenza [61].
There is evidence that cats, hamsters
and guinea pigs may naturally be infected
with CPIV/SV5 or a very closely
related virus [81, 144]. In addition, a
CPIV/SV5-like strain, termed SER, was
recently isolated from the lung of a fetus
of a breeding sow with porcine respiratory
and reproductive syndrome [79, 155].
Antibodies to CPVI have been demonstrated
in 20 of 44 wildlife species in eight
African countries [74]. Also, antibodies to
CPVI have been detected in non-captive
black bears (Ursus americanus) and fishers
(Martes pennanti) in Canada [120], suggesting
circulation of CPIV-like viruses in
wildlife animals. Even more interestingly,
CPIV/SV5-like viruses may infect humans
and non-human primates [39].
CPIV infection is usually restricted to
the upper respiratory tract in dogs of
two weeks of age or older [6]. Although
viremia is considered an uncommon event,
CPIV has been recovered from the lungs,
spleen, kidneys and liver of laboratory
dogs with mixed infections [22]. After experimental
infection of dogs, CPIV replicates
in cells of the nasal mucosa, pharynx,
trachea and bronchi. Small amounts
of virus can be recovered from the local
lymph nodes, but not from other lymphatic
tissues. In naturally infected dogs, simultaneous
infections with other viral and bacterial
agents are quite common and clinical
signs may be more severe [2, 22, 137].
Symptoms generally occur 2–8 days after
infection. CPIV produces mild symptoms
lasting less than six days, but

362 C. Buonavoglia, V. Martella
infection is usually complicated by other
pathogens. In the non-complicated forms,
clinical signs include low-grade rise in
temperature, deep sounding dry cough, watery
nasal discharge, pharyngitis and tonsillitis
[6]. Most dogs appear healthy and
active. In the complicated forms, described
mostly in immunocompromised animals
or young unvaccinated puppies, the symptoms
may progress and include lethargy,
fever, inappetance, and pneumonia.
CPIV has also been isolated from a dog
with temporary posterior paralysis [63]
and this isolate, termed CPI+, caused
acute encephalitis when injected intracranially
into gnotobiotic dogs [13]. From one
such experimentally infected dog, a variant,
termed CPI2, was isolated that had
phenotypic and genotypic differences from
CPI+. CPI2isattenuatedinferretsandit
more readily establishes persistent infections
in tissue culture cells. The biological
changes and the ability to block IFN
signaling have been mapped to the P/V-
N-terminal common domain of the V protein
[14–17, 38, 148].
In experimentally infected dogs petechial
hemorrhages have been described
in lung lobes between 3 and 8 days
post infection [2, 26]. Histological examination
has revealed catarrhal rhinitis
and tracheitis with mono- and polymorphonuclear
cell infiltrates in the mucosa
and submucosa. Bronchi and bronchioli
may contain leukocytes and cellular
debris.
Laboratory diagnosis may rely on viral
isolation from nasopharyngeal or laryngeal
swabs, using primary cells or cell lines
derived from dog kidneys. A wide range
of canine, feline, bovine, simian, and human
cells are permissive for CIPV and
monkey kidney cells have also been used
successfully [6]. In the first passage, the
virus usually does not induce cytopathic
effects and virus antigens may be demonstrated
by hemadsorption or immunofluorescence
[2, 44]. RT-PCR may also be
applied to respiratory secretions, nasopharyngeal/laryngeal
swabs and tracheal/lung
tissues [61]. Serological investigations by
hemagglutination inhibition and the virus
neutralization test may be useful to screen
animals for the presence of specific antibodies.
Attenuated vaccines have been developed
against CPIV. A parenteral CPIV
vaccine is available in combination with
other antigens. These vaccines alone rarely
provide protection against contracting the
infection, although they help to reduce the
severity of the disease. Vaccination of all
animals, notably of puppies, is indicated
in kennels or in pet shops. Strict hygiene
with thorough cleaning and disinfection
of cages and food and water containers,
good ventilation and adequate population
density are essential for controlling virus
spread.
6. CANINE REOVIRUS
Mammalian orthoreoviruses (MRV) are
non-enveloped, double-stranded (ds) RNA
viruses included in the genus Orthoreovirus
within the family Reoviridae. MRV
are responsible for either symptomatic or
asymptomatic infections in mammals and
possess a broad host range [157].
The reovirus genome contains ten
dsRNA segments, which are designed as
large (L, three segments), medium (M,
three segments), or small (S , four segments)
on the basis of the electrophoretic
mobility [118]. Three MRV serotypes have
been recognized by cross-evaluation with
specific sera in neutralization and inhibition
of haemagglutination assays [136,
141]. Neutralization and HA activities are
restricted to a single reovirus gene segment,
S 1 [165], that encodes for the proteins
σ1 andσ1s. The σ1 protein, a fibrous
trimer located on the outer capsid
of the virion [68, 69], is responsible for
viral attachment on cellular receptors [95,
167], serotype-specific neutralization [12],
and hemagglutination [166]. Analysis of

the S 1 gene of MRV belonging to different
serotypes has shown a strict correlation
between sequence similarity and
viral serotype [34, 56, 117]. Conversely,
the other genome segments do not display
any correlation to viral serotype, suggesting
that MRV have evolved independently
of serotype in the various species [25, 37,
71, 87, 94].
MRV have a wide geographic distribution
and can virtually infect all mammals,
including humans [157]. In carnivores,
MRV infections have been reported
sporadically, although all three serotypes
have been isolated from dogs and cats [21,
46, 51, 57, 89, 97, 102, 108, 113, 145].
As in other mammalians, the aetiological
role of MRV in respiratory diseases of
dogs is still unclear. MRV-1 strains have
been recovered from dogs with pneumonia
[97] or enteritis [4], in association with
either canine distemper virus or canine parvovirus
type 2. MRV-2 and MRV-3 have
been isolated from dogs with disease of the
upper respiratory tract [21] and with diarrhea
[51, 89], respectively. Only an MRV-3
enteric strain has been characterized at the
molecular level in the S1andL1segments.
The highest nucleotide identity was found
to a murine strain in the S 1 segment (93%)
and to human and bovine strains in the
L1 segments (90%), revealing the lack of
species-specific patterns [51]. By PCR, reovirus
RNA was detected in 50/192 rectal
swabs from dogs with diarrhea. Also, reovirus
RNA was detected in 9/12 ocular
swabs, in 10/19 nasal swabs of dogs with
ocular/nasal discharge, whereas it was not
detected in the oro-pharynx [57]. These
data suggest that reoviruses are common
in dogs, both in the enteric or in the respiratory
tract, although the viruses are shed
in low amounts. Experimental infections
with MRV in germ- and disease-free dogs
failed to give conclusive results [4, 80].
Accordingly, it appears that MRV do not
exert direct pathogenic activity and, more
likely, act in synergism with other respira-
Canine respiratory viruses 363
tory pathogens, aggravating the course of
concomitant infections [4].
Diagnosis of reovirus infection is usually
based on virus isolation on cell
cultures, electron microscopy and polyacrilamide
gel electrophoresis (PAGE).
These methods proved to be poorly sensitive
[115] and likely underestimate the
presence of MRV in animals and humans.
RT PCR protocols have been developed for
detection of MRV and for prediction of the
MRV serotype [51, 94, 115].
7. CANINE RESPIRATORY
CORONAVIRUS
Members of the Coronaviridae family
are enveloped viruses, 80–160 nm
in diameter, containing a linear positivestranded
RNA genome. Coronaviruses
are currently classified into four distinct
groups based on sequence analysis and
genome structure and on the antigenic
relationships [101, 139, 150]. The coronavirus
structural proteins include the
spike glycoprotein, the membrane glycoprotein
and the nucleocapsid protein.
The hemagglutinin-esterase glycoprotein
is found only on the surface of group 2
coronaviruses [60].
Three different coronaviruses have been
identified in dogs thus far [60, 125]. The
enteric canine coronaviruses (CCoV) are
distinguished into two genotypes, I and II,
and are included in group 1 coronaviruses
along with feline coronaviruses (FCoV)
type I and type II, transmissible gastroenteritis
virus of swine (TGEV), porcine
respiratory coronavirus (PRCoV), porcine
epidemic diarrhea virus (PEDV) and human
coronavirus 229E [59]. The evolution
of CCoV is tightly intermingled with that
of FCoV I and II [125]. Canine respiratory
coronavirus (CRCoV) was first detected
in the United Kingdom in 2003 from trachea
and lung tissues of dogs [60]. By
phylogenetic analysis of the polymerase,
CRCoV was found to segregate with group

364 C. Buonavoglia, V. Martella
2 coronaviruses, along with bovine coronaviruses
(BCoV) and human coronavirus
strain OC43 (HCV-OC43) [60]. Sequence
analysis of the S protein-encoding gene
revealed a high genetic similarity to the
bovine strain BCoV and to the human
strain OC43 (96.9 and 97.1% at the nucleotide
level and 96.0 and 95.2% at
the aa level, respectively) [60], suggesting
a recent common ancestor for the
three viruses and demonstrating the occurrence
of repeated host-species shifts [161].
Conversely, CRCoV was found to be genetically
and antigenically different from
the enteric canine coronaviruses (less than
21.2% aa in the S gene).
By RT-PCR, CRCoV RNA has been detected
both in asymptomatic and in symptomatic
dogs, that suffered from mild or
moderate respiratory disease [60]. Analysis
of archival samples has identified CR-
CoV in 2 out of 126 dogs affected by
respiratory diseases in Canada [58].
Taking advantage of the close genetic
relatedness between CCRoV and BCoV,
ELISA assays have been set up using
BCoV antigen and have been used to
screen canine sera, revealing the presence
of specific antibodies in 30.1% of dogs at
the time of entry in a rehoming kennel [60].
In a further study, antibodies were detected
in 22.2% and 54.2% of dogs on the
day of entry into working kennels in London
and Warwickshire, respectively [62].
In larger sero-epidemiological surveys, the
prevalence of CRCoV was demonstrated
to be 54.7% in North America, 36.6%
in the United Kingdom [126], 17.8% in
Japan [84] and 32% in Italy [53] while
there was no evidence for CRCoV-specific
antibodies in cats [84]. By examining
the relationship between the age of dogs
and the presence of CRCoV antibodies, a
steady increase in the seropositivity rates
was observed, with the highest prevalence
among dogs of 7–8 years (68.4%) [126].
Attempts to isolate CRCoV from tissue
of the respiratory tract, using canine
lung fibroblasts, MDCK, HRT-18G and
fewf-4 cells were unsuccessful [60,84] and
this has hampered, thus far, the evaluation
of CRCoV patho-biological properties and
the CRCoV role in canine respiratory diseases.
The role of CRCoV in ITB is not
clear. Sero-conversion was observed in
immunologically-naïve dogs after introduction
in a kennel where infected dogs
were housed, revealing a highly contagious
nature [60]. Dogs sero-negative to CRCoV
were statistically more prone to develop
respiratory disease than dogs with antibodies
to CRCoV, providing indirect evidence
for a pathogenic role of CRCoV [60]. It
is likely that CRCoV alone may induce
only subclinical or mild respiratory symptoms.
However, coronavirus replication
can damage the respiratory epithelium
and lead to bacterial superinfections. The
human respiratory coronavirus 229E can
disrupt the respiratory epithelium and
cause ciliary dyskinesia [40]. Accordingly,
virus-induced alterations of the respiratory
epithelium would trigger the replication of
other pathogens, causing respiratory diseases
resembling the ITB complex.
8. PANTROPIC CANINE
CORONAVIRUS
Enteric CCoV usually cause mild to
severe diarrhea in pups, whereas fatal
infections have been associated mainly
with concurrent infections by canine parvovirus,
canine adenovirus type 1 or canine
distemper virus [50, 123, 124].
Thus far, two genotypes of enteric
CCoV have been described, namely CCoV
type I and CCoV type II [125]. Molecular
methods able to distinguish between the
two genotypes have revealed that mixed infections
by both genotypes occur at high
frequency in dogs [52].
Recently, a fatal, systemic disease
caused by a highly virulent CCoV strain

was reported, which was characterized
by severe gastrointestinal and respiratory
symptoms [29]. The disease occurred in
seven dogs housed in a pet shop in the
Apulia region, Italy. The dogs displayed
fever (39.5–40 ◦ C), lethargy, inappetance,
respiratory distress, vomiting, hemorrhagic
diarrhea, and neurological signs (ataxia,
seizures) followed by death within 2 days
after the onset of the symptoms. A marked
leukopenia, with total WBC counts below
50% of the baseline values, was also
reported. Necropsy examination revealed
severe gross lesions in the tonsils, lungs,
liver, spleen and kidneys. Extensive lobar
subacute bronchopneumonia was evidenced
both in the cranial and caudal
lobes, along with effusions in the thoracic
cavity.
By genotype-specific real-time RT-PCR
assays, CCoV type II RNA was detected in
the intestinal content and parenchymatous
organs, including the lungs, and a coronavirus
strain was successfully isolated on
cell cultures from lungs and other tissues.
Sequence analysis of the 3’ end of the
viral genome showed a point mutation in
the S protein and a truncated form of the
nonstructural protein 3b, due to the presence
of a 38-nt deletion and to a frame shift
in the sequence downstream of the deletion
that introduced an early stop codon
in ORF3b. Either point mutations or deletions
in the structural spike glycoprotein
and in the nonstructural proteins have been
associated to changes in tropism and virulence
of coronaviruses [72, 78, 83, 93,
160]. The porcine respiratory coronavirus
(PRCoV), a spike (S) gene deletion mutant
of transmissible gastroenteritis virus
(TGEV), causes mild or subclinical respiratory
infections in pigs [93].
Experimental infection of dogs with the
virus isolate resulted in a severe systemic
disease that mimicked the clinical signs
observed in the outbreak. However, older
puppies were able to recover from the
infection. The pathogenic CCoV variants
Canine respiratory viruses 365
should be suspected when unexplainable
episodes of severe to fatal disease occur in
pups. Epidemiological studies are required
to determine whether the pantropic CCoV
strain is a new coronavirus variant emerging
in the canine population or if it is a
wide-spread infectious agent of dogs that
usually goes undetected. Vaccination trials
are necessary to determine whether the
CCoV vaccines currently available are effective
against the highly virulent CCoV
strain.
9. CONCLUSIONS
The development of new diagnostic
techniques and the extensive use of molecular
analysis are quickly providing an
increasing amount of information on the
epidemiology of respiratory viruses, on the
molecular basis of pathogenicity and on
the mechanisms that drive virus evolution.
In the last decades, evidence has been collected
for the emergence of novel viruses
by host species shift or by change of tissue
tropism due to genome mutations. Prophylaxis
of the ITB complex relies on the use
of vaccines based on selected pathogens
(CAV-2, CPIV and Bordetella bonchiseptica)
and those vaccines are not always
effective in preventing ITB, suggesting that
other pathogens may also play a role in
respiratory diseases of dogs. Whether the
detection of new respiratory pathogens requires
the development of novel prophylaxis
tools is an issue that surely deserves
more attention. At the same time, intensification
of surveillance activity is paramount
to monitor the emergence and spread of
novel pathogens, to investigate their epidemiology
and plan adequate measures of
control.
REFERENCES
[1] Ajiki M., Takamura K., Hiramatsu K.,
Nakai M., Sasaki N., Konishi S., Isolation

366 C. Buonavoglia, V. Martella
and characterization of parainfluenza 5
virus from a dog, Nippon Juigaku Zasshi
(1982) 44:607–618.
[2] Appel M., Percy D.H., SV5-like parainfluenza
virus in dogs, J. Am. Vet. Med.
Assoc. (1970) 156:1778–1781.
[3] Appel M., Bistner S.I., Menegus M., Albert
D.A., Carmichael L.E., Pathogenicity of
low-virulence strains of two canine adenoviruses,
Am. J. Vet. Res. (1973) 34:543–
550.
[4] Appel M., Reovirus, in: Appel M.J. (Ed.),
Virus infections of carnivores, Elsevier
Science Publisher, Amsterdam, 1987, pp.
95–96.
[5] Appel M., Canine adenovirus type 2
(Infectious Laryngotracheitis Virus), in:
Appel M.J. (Ed.), Virus Infections of
Carnivores, Elsevier Science Publisher,
Amsterdam, 1987, pp. 45–51.
[6] Appel M., Binn L.N., Canine infectious
tracheobronchitis. Short review: kennel
cough, in: Appel M.J. (Ed.), Virus infections
of carnivores, Elsevier Science
Publisher, Amsterdam, 1987, pp. 201–211.
[7] Appel M.J., Menegus M., Parsonson I.M.,
Carmichael L.E., Pathogenesis of canine
herpesvirus in specific-pathogen-free dogs:
5- to 12-week-old pups, Am. J. Vet. Res.
(1969) 30:2067–2073.
[8] Appel M.J.G., Carmichael L.E., Robson
D.S., Canine adenovirus type 2-induced immunity
to two canine adenoviruses in pups
with maternal antibody, Am. J. Vet. Res.
(1975) 36:1199–1202.
[9] Appel M.J., Canine infectious tracheobronchitis
(kennel cough): a status report,
Compend. Contin. Educ. Pract. Vet. (1981)
3:70–79.
[10] Assaf R., Marsolais G., Yelle J., Hamelin
C., Unambiguous typing of canine adenovirus
isolates by deoxyribonucleic acid
restriction-endonuclease analysis, Can. J.
Comp. Med. (1983) 47:460–463.
[11] Azetaka M., Konishi S., Kennel cough
complex: confirmation and analysis of the
outbreak in Japan, Nippon Juigaku Zasshi
(1988) 50:851–858.
[12] Bassel-Duby R., Spriggs D.R., Tyler K.L.,
Fields B.N., Identification of attenuating
mutations on the reovirus type 3
S1 double-stranded RNA segment with a
rapid sequencing technique, J. Virol. (1986)
60:64–67.
[13] Baumgärtner W.K., Metzler A.E.,
Krakowka S., Koestner A., In vitro
identification and characterization of a
virus isolated from a dog with neurological
dysfunction, Infect. Immun. (1981)
31:1177–1183.
[14] Baumgärtner W.K., Krakowka S., Koestner
A., Evermann J., Acute encephalitis and
hydrocephalus in dogs caused by canine
parainfluenza virus, Vet. Pathol. (1982)
19:79–92.
[15] Baumgärtner W., Krakowka S., Blakeslee
J., Evolution of in vitro persistence of two
strains of canine parainfluenza virus. Brief
report, Arch. Virol. (1987) 93:147–154.
[16] Baumgärtner W., Krakowka S., Blakeslee
J.R., Persistent infection of Vero cells
by paramyxoviruses. A morphological and
immunoelectron microscopic investigation,
Intervirology (1987) 27:218–223.
[17] Baumgärtner W., Krakowka S., Durchfeld
B., In vitro cytopathogenicity and in vivo
virulence of two strains of canine parainfluenza
virus, Vet. Pathol. (1991) 28:324–
331.
[18] Bemis D.A., Carmichael L.E., Appel
M.J.G., Naturally occurring respiratory disease
in a kennel caused by Bordetella bronchiseptica,
Cornell Vet. (1977) 67:282–293.
[19] Benetka V., Weissenbock H., Kudielka I.,
Pallan C., Rothmuller G., Mostl K., Canine
adenovirus type 2 infection in four puppies
with neurological signs, Vet. Rec. (2006)
158:91–94.
[20] Binn L.N., Lazar E.C., Rogul M., Shepler
V.M., Swango L.J., Claypoole T., Hubbard
D.W., Asbill S.G., Alexander A.D., Upper
respiratory disease in mility dogs: bacterial
mycoplasma and viral studies, Am. J. Vet.
Res. (1968) 29:1809–1815.
[21] Binn L.N., Marchwicki R.H., Keenan K.P.,
Strano A.J., Engler W.R., Recovery of reovirus
type 2 from an immature dog with
respiratory tract disease, Am. J. Vet. Res.
(1977) 38:927–929.
[22] Binn L.N., Alford J.P., Marchwicki R.H.,
Keefe T.J., Beattie R.J., Wall H.G., Studies
of respiratory disease in random-source
laboratory dogs: viral infections in unconditioned
dogs, Lab. Anim. Sci. (1979)
29:48–52.
[23] Biron C.A., Sen G.C., Interferons and
other cytokines, in: Knipe D.M., Howley
P.M. (Eds.), Fields Virology, 4th edition,
Lippincott Williams & Wilkins,
Philadelphia, 2001, pp. 321–351.
[24] Bittle J.L., Grant W.A., Scott F.W., Canine
and feline immunization guidelines in

1982, J. Am. Vet. Med. Assoc. (1982)
181:332–335.
[25] Breun L.A., Broering T.J., McCutcheon
A.M., Harrison S.J., Luongo C.L., Nibert
M.L., Mammalian reovirus L2 gene and l2
core spike protein sequences and wholegenome
comparison of reoviruses type 1
Lang, type 2 Jones, and type 3 Dearing,
Virology (2001) 287:333–348.
[26] Brown A.L., Bihr J.G., Vitamvas J.A.,
Miers L., An alternative method for evaluating
potency of modified live canine parainfluenza
virus vaccine, J. Biol. Stand. (1978)
6:271–281.
[27] Brown I.H., The epidemiology and evolution
of influenza viruses in pigs, Vet.
Microbiol. (2000) 74:29–46.
[28] Buonavoglia C., Sala V., Indagine sierologica
nei cani sulla presenza di anticorpi
verso ceppi di virus influenzali umani tipo
A, Clin. Vet. (1983) 106:81–83.
[29] Buonavoglia C., Decaro N., Martella V.,
Elia G., Campolo M., Desario C., Tempesta
M., Canine coronavirus highly pathogenic
for dogs, Emerg. Infect. Dis. (2006)
12:492–494.
[30] Burek K.A., Gulland F.M., Sheffield G.,
Beckmen K.B., Keyes E., Spraker T.R.,
Smith A.W., Skilling D.E., Evermann
J.F., Stott J.L., Saliki J.T., Trites A.W.,
Infectious disease and the decline of Steller
sea lions (Eumetopias jubatus) inAlaska,
USA: insights from serologic data, J. Wildl.
Dis. (2005) 41:512–524.
[31] Burr P.D., Campbell M.E., Nicolson L.,
Onions D.E., Detection of canine herpesvirus
1 in a wide range of tissues using
the polymerase chain reaction, Vet.
Microbiol. (1996) 53:227–237.
[32] Carmichael L.E., Squire R.A., Krook L.,
Clinical and pathologic features of a fatal
viral disease of newborn pups, Am. J. Vet.
Res. (1965) 26:803–814.
[33] Carmichael L.E., Greene C.E., Canine herpesvirus
infection, in: Greene C.E. (Ed.),
Infectious diseases of the dog and cat,
WB Saunders Co, Philadelphia, 1998,
pp. 28–32.
[34] Cashdollar L.W., Chmelo R.A., Wiener
J.R., Joklik W.K., The sequence of the
S1 genes of the three serotypes of reovirus,
Proc. Natl. Acad. Sci. USA (1985)
82:24–28.
[35] Castleman W.L., Bronchiolitis obliterans
and pneumonia-induced in young dogs by
Canine respiratory viruses 367
experimental adenovirus infection, Am. J.
Pathol. (1985) 119:495–504.
[36] Chalker V.J., Brooks H.W., Brownlie J.,
The association of Streptococcus equi
subsp. zooepidemicus with canine infectious
respiratory disease, Vet. Microbiol.
(2003) 95:149–156.
[37] Chappel J.D., Goral M.I., Rodgers S.E., de-
Pamphilis C.W., Dermody T.S., Sequence
diversity within the reovirus S2 gene: reovirus
genes reassort in nature, and their
termini are predicted to form a panhandle
motif, J. Virol. (1994) 68:750–756.
[38] Chatziandreou N., Young D., Andrejeva J.,
Goodbourn S., Randall R.E., Differences in
interferon sensitivity and biological properties
of two related isolates of simian virus
5: a model for virus persistence, Virology
(2002) 293:234–242.
[39] Chatziandreou N., Stock N., Young D.,
Andrejeva J., Hagmaier K., McGeoch D.J.,
Randall R.E., Relationships and host range
of human, canine, simian and porcine isolates
of simian virus 5 (parainfluenza virus
5), J. Gen. Virol. (2004) 85:3007–3016.
[40] Chilvers M.A., McKean M., Rutman A.,
Myint B.S., Silverman M., O’Callaghan C.,
The effects of coronavirus on human nasal
ciliated respiratory epithelium, Eur. Respir.
J. (2001) 18:965–970.
[41] Claas E.C., Osterhaus A.D., van Beek R.,
De Jong J.C., Rimmelzwaan G.F., Senne
D.A., Krauss S., Shortridge K.F., Webster
R.G., Human influenza A H5N1 virus related
to a highly pathogenic avian influenza
virus, Lancet (1998) 351:472–477.
[42] Cornwell H.J., McCandlish I.A., Thompson
H., Laird H.M., Wright N.G., Isolation
of parainfluenza virus SV5 from dogs
with respiratory disease, Vet. Rec. (1976)
98:301–302.
[43] Cornwell H.J., Koptopoulos G., Thompson
H., McCandlish I.A., Wright N.G.,
Immunity to canine adenovirus respiratory
disease: a comparison of attenuated CAV-1
and CAV-2 vaccines, Vet. Rec. (1982)
110:27–32.
[44] Crandell R.A., Brumlow W.B., Davison
V.E., Isolation of a parainfluenza virus from
sentry dogs with upper respiratory disease,
Am. J. Vet. Res. (1968) 29:2141–2147.
[45] Crawford P.C., Dubovi E.J., Castleman
W.L., Stephenson I., Gibbs E.P., Chen L.,
Smith C., Hill R.C., Ferro P., Pompey J.,
Bright R.A., Medina M.J., Johnson C.M.,
Olsen C.W., Cox N.J., Klimov A.I., Katz

368 C. Buonavoglia, V. Martella
J.M., Donis R.O., Transmission of equine
influenza virus to dogs, Science (2005)
310:482–485.
[46] Csiza C.K., Characterization and serotyping
of three feline reovirus isolates, Infect.
Immun. (1974) 9:159–166.
[47] Curtis R., Jemmet J.E., Furminger I.G.S.,
The pathogenicity of an attenuated strain
of canine adenovirus type 2 (CAV-2), Vet.
Rec. (1978) 103:380-381.
[48] Curtis R., Barnett K.C., The ’blue eye’ phenomenon,
Vet. Rec. (1983) 112:347–353.
[49] Davison A.J., Benko M., Harrach B.,
Genetic content and evolution of adenoviruses,
J. Gen. Virol. (2003) 84:2895–
2908.
[50] Decaro N., Camero M., Greco G., Zizzo
N., Elia G., Campolo M., Pratelli A.,
Buonavoglia C., Canine distemper and related
diseases: report of a severe outbreak
in a kennel, New Microbiol. (2004) 27:177–
181.
[51] Decaro N., Campolo M., Desario C., Ricci
D., Camero M., Lorusso E., Elia G.,
Lavazza A., Martella V., Buonavoglia C.,
Virological and molecular characterization
of a Mammalian orthoreovirus type 3 strain
isolated from a dog in Italy, Vet. Microbiol.
(2005) 109:19–27.
[52] Decaro N., Martella V., Ricci D., Elia G.,
Desario C., Campolo M., Cavaliere N., Di
Trani L., Tempesta M., Buonavoglia C.,
Genotype-specific fluorogenic RT-PCR assays
for the detection and quantitation of
canine coronavirus type I and type II RNA
in faecal samples of dogs, J. Virol. Methods
(2005) 130:72–78.
[53] Decaro N., Desario C., Elia G., Mari V.,
Lucente M.S., Cordioli P., Colaianni M.L.
Martella V., Buonavoglia C., Serological
and molecular evidence that canine respiratory
coronavirus is circulating in Italy, Vet.
Microbiol. (2006) (in press).
[54] Didcock L., Young D.F., Goodbourn S.,
Randall R.E., Sendai virus and simian virus
5 block activation of interferon-responsive
genes: importance for virus pathogenesis, J.
Virol. (1999) 73:3125–3133.
[55] Ditchfield J., MacPherson L.W., Zbitnew
A., Association of a canine adenovirus
(Toronto A26/61) with an outbreak of
laryngotracheitis (kennel cough). A preliminary
report, Can. Vet. J. (1962) 3:238–247.
[56] Duncan R., Horne D., Cashdollar L.W.,
Joklik W.K., Lee P.W.K., Identification of
conserved domains in the cell attachment
proteins of the three serotypes of reovirus,
Virology (1990) 174:339–409.
[57] Elia G., Lucente M.S., Bellacicco A.L.,
Camero M., Cavaliere N., Decaro N.,
Martella V., Assessment of reovirus epidemiology
in dogs, in: Proc. 16th Eur.
Congress of Clinical Microbiology and
Infectious Diseases, Nice, 2006, p. 1674.
[58] Ellis J.A., McLean N., Hupaelo R., Haines
D.M., Detection of coronavirus in cases
of tracheobronchitis in dogs: a retrospective
study from 1971 to 2003, Can. Vet. J.
(2005) 46:447–448.
[59] Enjuanes L., Brian D., Cavanagh D.,
Holmes K., Lai M.M.C., Laude H., Masters
P., Rottier P., Siddell S., Spaan W.J.M.,
Taguchi F., Talbot P., Coronaviridae, in:
van Regenmortel M.H.V., Fauquet C.M.,
Bishop D.H.L., Carstens E.B., Estes M.K.,
Lemon S.M., et al. (Eds.), Virus Taxonomy,
Classification and Nomenclature of
Viruses, Academic Press, New York, 2000,
pp. 835–849.
[60] Erles K., Toomey C., Brooks H.W.,
Brownlie J., Detection of a group 2 coronavirus
in dogs with canine infectious respiratory
disease, Virology (2003) 310:216–
223.
[61] Erles K., Dubovi E.J., Brooks H.W.,
Brownlie J., Longitudinal study of viruses
associated with canine infectious respiratory
disease, J. Clin. Microbiol. (2004)
42:4524–4529.
[62] Erles K., Brownlie J., Investigation into
the causes of canine infectious respiratory
disease: antibody responses to canine respiratory
coronavirus and canine herpesvirus
in two kennelled dog populations, Arch.
Virol. (2005) 150:1493–1504.
[63] Evermann J.F., Lincoln J.D., McKiernan
A.J., Isolation of a paramyxovirus from the
cerebrospinal fluid of a dog with posterior
paresis, J. Am. Vet. Med. Assoc. (1980)
177:1132–1134.
[64] Evermann J.F., LeaMaster B.R., McElwain
T.F., Potter K.A., McKeirnan A.J., Green
J.S., Natural infection of captive coyote
pups with a herpesvirus antigenically related
to canine herpesvirus, J. Am. Vet.
Med. Assoc. (1984) 185:1288–1290.
[65] Fairchild G.A., Cohen D., Serologic study
of canine adenovirus (Toronto A26/61) infection
in dogs, Am. J. Vet. Res. (1969)
30:923–928.
[66] Fouchier R.A., Schneeberger P.M.,
Rozendaal F.W., Broekman J.M., Kemink

S.A., Munster V., Kuiken T., Rimmelzwaan
G.F., Schutten M., Van Doornum G.J.,
Koch G., Bosman A., Koopmans M.,
Osterhaus A.D., Avian influenza A virus
(H7N7) associated with human conjunctivitis
and a fatal case of acute respiratory
distress syndrome, Proc. Natl. Acad. Sci.
USA (2004) 101:1356–1361.
[67] Fouchier R.A., Munster V., Wallensten
A., Bestebroer T.M., Herfst S., Smith D.,
Rimmelzwaan G.F., Olsen B., Osterhaus
A.D., Characterization of a novel influenza
A virus hemagglutinin subtype (H16) obtained
from black-headed gulls, J. Virol.
(2005) 79:2814–2822.
[68] Fraser R.D., Furlong D.B., Trus B.L.,
Nibert M.L., Fields B.N., Steven
A.C., Molecular structure of the cellattachment
protein of reovirus: correlation
of computer-processed electron micrographs
with sequence-based predictions, J.
Virol. (1990) 64:2990-3000.
[69] Furlong D.B., Nibert M.L., Fields B.N.,
Sigma 1 protein of mammalian reoviruses
extends from the surfaces of viral particles,
J. Virol. (1988) 62:246-256.
[70] Garcelon D.K., Wayne R.K., Gonzales
B.J., A serologic survey of the island fox
(Urocyon littoralis) on the Channel Islands,
California, J. Wildl. Dis. (1992) 28:223–
229.
[71] Goral M.I., Mochow-Grundy M., Dermody
T.S., Sequence diversity within the reovirus
S3 gene: reoviruses evolve independently
of host species, geographic locale, and date
of isolation, Virology (1996) 216:265–271.
[72] Guan Y., Zheng B.J., He Y.Q., Liu X.L.,
Zhuang Z.X., Cheung C.L., et al., Isolation
and characterization of viruses related to
the SARS coronavirus from animals in
southern China, Science (2003) 302:276–
278.
[73] Haanes E.J., Tomlinson C.C., Genomic organization
of the canine herpesvirus US
region, Virus Res. (1998) 53:151–162.
[74] Hamblin C., Hedger R.S., Neutralising antibodies
to parainfluenza 3 virus in African
wildlife, with special reference to the Cape
buffalo (Syncerus caffer), J. Wildl. Dis.
(1978) 14:378-388.
[75] Hamelin C., Marsolais G., Assaf R.,
Interspecific differences between the DNA
restriction profiles of canine adenoviruses,
Experientia (1984) 40:482.
[76] Hamelin C., Jouvenne P., Assaf R.,
Association of a type-2 canine adenovirus
with an outbreak of diarrhoeal
Canine respiratory viruses 369
disease among a large dog congregation, J.
Diarrhoeal Dis. Res. (1985) 3:84–87.
[77] Hashimoto A., Hirai K., Yamaguchi T.,
Fujimoto Y., Experimental transplacental
infection of pregnant dogs with canine herpesvirus,
Am. J. Vet. Res. (1982) 43:844–
850.
[78] Haspel M.V., Lampert P.W., Oldstone M.B.,
Temperature-sensitive mutants of mouse
hepatitis virus produce a high incidence of
demyelination, Proc. Natl. Acad. Sci. USA
(1978) 75:4033–4036.
[79] Heinen E., Herbst W., Schmeer N.,
Isolation of a cytopathogenic virus from a
case of porcine reproductive and respiratory
syndrome (PRRS) and its characterization
as parainfluenza virus type 2, Arch. Virol.
(1998) 143:2233–2239.
[80] Holzinger E.A., Griesemer R.A., Effects of
reovirus type 1 on germfree and diseasefree
dogs, Am. J. Epidemiol. (1966)
84:426–430.
[81] Hsiung G.D., Parainfluenza-5 virus.
Infection of man and animal, Prog. Med.
Virol. (1972) 14:241–274.
[82] Hu R.L., Huang G., Qiu W., Zhong Z.H.,
Xia X.Z., Yin Z., Detection and differentiation
of CAV-1 and CAV-2 by polymerase
chain reaction, Vet. Res. Commun. (2001)
25:77–84.
[83] Jonassen C.M., Kofstad T., Larsen I.L.,
Lovland A., Handeland K., Follestad A.,
et al., Molecular identification and characterization
of novel coronaviruses infecting
graylag geese (Anser anser), feral
pigeons (Columbia livia) and mallards
(Anas platyrhynchos), J. Gen. Virol. (2005)
86:1597–1607.
[84] Kaneshima T., Hohdatsu T., Satoh K.,
Takano T., Motokawa K., Koyama H., The
prevalence of a group 2 coronavirus in dogs
in Japan, J. Vet. Med. Sci. (2006) 68:21–25.
[85] Karpas A., Garcia F.G., Calvo F., Cross
R.E. Experimental production of canine tracheobronchitis
(kennel cough) with canine
herpesvirus isolated from naturally infected
dogs, Am. J. Vet. Res. (1968) 29:1251–
1257.
[86] Karpas A., King N.W., Garcia F.G., Calvo
F., Cross R.E., Canine tracheobronchitis;
Isolation and characterization of the agent
with experimental reproduction of the disease,
Proc. Soc. Exp. Biol. Med. (1968)
127:45–52.
[87] Kedl R., Schmechel S., Schiff L.,
Comparative sequence analysis of the

370 C. Buonavoglia, V. Martella
reovirus S4 genes from 13 serotype 1 and
serotype 3 field isolates, J. Virol. (1995)
69:552–559.
[88] Kimber K.R., Kollias G.V., Dubovi E.J.,
Serologic survey of selected viral agents
in recently captured wild North American
river otters (Lontra canadensis), J. Zoo
Wildl. Med. (2000) 31:168–175.
[89] Kokubu T., Takahashi T., Takamura K.,
Yasuda H., Hiramatsu K., Nakai M.,
Isolation of a reovirus type 3 from dogs
with diarrhea, J. Vet. Med. Sci. (1993)
55:453–454.
[90] Koptopoulos G., Cornwell H.J.C., Canine
adenoviruses: a review, Vet. Bull. (1981)
51:135–142.
[91] Krakowka S., Canine herpesvirus-1, in:
Olsen R.G., Krakowka S., Blakeslee J.R. Jr.
(Eds.), Comparative pathobiology of viral
disease, CRC Press, Boca Raton, 1985, pp.
137–144.
[92] Lamb R.A., Kolakofsky D.,
Paramyxoviridae: The viruses and their
replication, in: Knipe D.M., Howley
P.M. (Eds.), Fields virology, 4th edition,
Lippincott Williams & Wilkins,
Philadelphia, 2001, pp. 1305–1340.
[93] Laude H., Van Reeth K., Pensaert M.,
Porcine respiratory coronavirus: molecular
features and virus-host interactions, Vet.
Res. (1993) 24:125–150.
[94] Leary P.L., Erker J.C., Chalmers M.L.,
Cruz A.T., Wetzel J.D., Desai S.M.,
Mushahwar I.K., Dermody T.S., Detection
of mammalian reovirus RNA by using reverse
transcription-PCR: sequence diversity
within the l3-encoding L1 gene, J. Clin.
Microbiol. (2002) 40:1368–1375.
[95] Lee P.W., Hayes E.C., Joklik W.K., Protein
s1 is the reovirus cell attachment protein,
Virology (1981) 108:156–163.
[96] Limbach K.J., Limbach M.P., Conte D.,
Paoletti E., Nucleotide sequenze of the
genes encoding the canine herpesvirus gB,
gC and gD homologues, J. Gen. Virol.
(1994) 75:2029–2039.
[97] Lou T.Y., Wenner H.A., Natural and experimental
infection of dogs with reovirus type
1: Pathogenecity of the strain for other animals,
Am. J. Hyg. (1963) 77:293–304.
[98] Macartney L., Cavanagh H.M., Spibey N.,
Isolation of canine adenovirus-2 from the
faeces of dogs with enteric disease and
its unambiguous typing by restriction endonuclease
mapping, Res. Vet. Sci. (1988)
44:9–14.
[99] Maeda K., Yokoyama N., Fujita K.,
Maejima M., Mikami T., Heparin-binding
activity of feline herpesvirus type 1 glycoproteins,
Virus Res. (1997) 52:169–176.
[100] Markwell M., New frontiers opened by
the exploration of host cell receptor, in:
Kingsbury D. (Ed.), The Paramyxoviruses,
Plenum Press, New York, 1991, pp. 407-
426.
[101] Marra M.A., Jones S.J., Astell C.R., Holt
R.A., et al., The Genome sequence of
the SARS-associated coronavirus, Science
(2003) 300:1399–1404.
[102] Marshall J.A., Kennett M.L., Rodger S.M.,
Studdert M.J., Thompson W.L., Gust I.D.,
Virus and virus-like particles in the faeces
of cats with and without diarrhea, Aust. Vet.
J. (1987) 64:100–105.
[103] Martina B.E., Harder T.C., Osterhaus A.D.,
Genetic characterization of the unique short
segment of phocid herpesvirus type 1 reveals
close relationships among alphaherpesviruses
of hosts of the order Carnivora,
J. Gen. Virol. (2003) 84:1427–1430.
[104] Marusyk R.G., Norrby E., Lundqvist U.,
Biophysical comparison of two canine adenoviruses,
J. Virol. (1970) 5:507–512.
[105] Marusyk R.G., Yamamoto T.,
Characterization of canine adenoviruse
hemagglutinin, Can. J. Microbiol. (1971)
17:151–155.
[106] Marusyk R.G., Hammarskjold M.L., The
genetic relationship of two canine adenoviruses
as determined by nucleic acid hybridization,
Microbios (1972) 5:259–264.
[107] Marusyk R.G., Comparison of the immunological
properties of two canine adenoviruses,
Can. J. Microbiol. (1972) 18:817–
823.
[108] Massie E.L., Shaw E.D., Reovirus type 1
in laboratory dogs, Am. J. Vet. Res. (1966)
27:783–787.
[109] Matthews R.E.F., Classification and
nomenclature of viruses, Fourth Report of
the International Committee on Taxonomy
of Viruses, Intervirology (1982) 17:4–199.
[110] McCandlish I.A., Thompson H., Cornwell
H.J., Wright N.G., A study of dogs with
kennel cough, Vet. Rec. (1978) 102:293–
301.
[111] Meng Q., Qiao J., Guo X., Cloning
and sequence analysis of fusion protein
gene of canine parainfluenza
virus wildtype strain, in: Gene bank,
http://www.ncbi.nlm.nih.gov, 2003.

[112] Miyoshi M., Ishii Y., Takiguchi M., Takada
A., Yasuda J., Hashimoto A., Okazaki K.,
Kida H., Detection of canine herpesvirus
DNA in the ganglionic neurons and the
lymph node lymphocytes of latently infected
dogs, J. Vet. Med. Sci. (1999)
61:375–379.
[113] Mochizuki M., Uchizono S., Experimental
infections of feline reovirus serotype 2 isolates,
J. Vet. Med. Sci. (1993) 55:469–470.
[114] Morrison M.D., Onions D.E., Nicolson L.,
Complete DNA sequence of canine adenovirus
type 1, J. Gen. Virol. (1997) 78:873–
878.
[115] Muscillo M., La Rosa G., Marianelli C.,
Zaniratti S., Capobianchi M.R., Cantiani L.,
Carducci A., A new RT-PCR method for
the identification of reoviruses in seawater
samples, Water Res. (2001) 35:548–556.
[116] Nakamichi K., Ohara K., Matsumoto Y.,
Otsuka H., Attachment and penetration
of canine herpesvirus 1 in non-permissive
cells, J. Vet. Med. Sci. (2000) 62:965–970.
[117] Nibert M.L., Dermody T.S., Fields B.N.,
Structure of the reovirus cell-attachment
protein: a model for the domain organization
of s1, J. Virol. (1990) 64:2976–2989.
[118] Nibert M.L., Schiff L.A., Reoviruses and
their replication, in: Knipe D.M., Howley
P.M. (Eds.), Fields Virology, 4th edition,
Lippincott Williams & Wilkins,
Philadelphia, 2001, pp. 1679–1728.
[119] Okuda Y., Ishida K., Hashimoto A.,
Yamaguchi T., Fukushi H., Hirai K.,
Carmichael L.E., Virus reactivation in
bitches with a medical history of herpesvirus
infection, Am. J. Vet. Res. (1993)
54:551–554.
[120] Philippa J.D.W., Leighton P.J., Nielsen O.,
Pagliarulo M., Schwantje H., Shury T.,
Van Herwijnen R., Martina B., Kuiken T.,
Van de Bildt M.W.G., Osterhaus A.D.M.E.,
Antibodies to selected pathogens in freeranging
terrestrial carnivores and marine
mammals in Canada, Vet. Rec. (2004)
155:135–140.
[121] Poole E., He B., Lamb R.A., Randall R.E.,
Goodbourn S., The V proteins of simian
virus 5 and other paramyxoviruses inhibit
induction of interferon-β, Virology (2002)
303:33–46.
[122] Poste G., Lecatsas G., Apostolov K.,
Electron microscope study of the morphogenesis
of a new canine herpesvirus in dog
kidney cells, Arch. Gesamte. Virusforsch.
(1972) 39:317–329.
Canine respiratory viruses 371
[123] Pratelli A., Tempesta M., Roperto F.P.,
Sagazio P., Carmichael L.E., Buonavoglia
C., Fatal coronavirus infection in puppies
following canine parvovirus 2b infection, J.
Vet. Diagn. Invest. (1999) 11:550–553.
[124] Pratelli A., Martella V., Elia G., Tempesta
M., Guarda F., Capucchio M.T., Carmichael
L.E., Buonavoglia C., Severe enteric disease
in an animal shelter associated with
dual infection by canine adenovirus type
1 and canine coronavirus, J. Vet. Med.
B Infect. Dis. Vet. Public Health (2001)
48:385–392.
[125] Pratelli A., Martella V., Decaro N., Tinelli
A., Camero M., Cirone F., Elia G., Cavalli
A., Corrente M., Greco G., Buonavoglia D.,
Gentile M., Tempesta M., Buonavoglia C.,
Genetic diversity of a canine coronavirus
detected in pups with diarrhoea in Italy, J.
Virol. Methods (2003) 10:9–17.
[126] Priestnall S.L., Brownlie J., Dubovi E.J.,
Erles K., Serological prevalence of canine
respiratory coronavirus, Vet. Microbiol.
(2006) 115:43–53.
[127] Randall R.E., Young D.F., Goswami K.K.,
Russell W.C., Isolation and characterization
of monoclonal antibodies to simian virus 5
and their use in revealing antigenic differences
between human, canine and simian
isolates, J. Gen. Virol. (1987) 68:2769–
2780.
[128] Randolph J.F., Moise N.S., Scarlett J.M.,
Shin S.J., Blue J.T., Bookbinder P.R.,
Prevalence of mycoplasmal and ureaplasmal
recovery from tracheobronchial
lavages and of mycoplasmal recovery from
pharyngeal swab specimens in cats with
or without pulmonary disease, Am. J. Vet.
Res. (1993) 24:897–900.
[129] Reading M.J., Field H.J., Detection of high
levels of canine herpes virus-1 neutralising
antibody in kennel dogs using a novel
serum neutralisation test, Res. Vet. Sci.
(1999) 66:273-275.
[130] Rémond M., Sheldrick P., Lebreton F.,
Nardeux P., Foulon T., Gene organization
in the UL region and inverted repeats of the
canine herpesvirus genome, J. Gen. Virol.
(1996) 77:37–48.
[131] Reubel G.H., Pekin J., Venables D., Wright
J., Zabar S., Leslie K., Rothwell T.L., Hinds
L.A., Braid A., Experimental infection of
European red foxes (Vulpes vulpes) with
canine herpesvirus, Vet. Microbiol. (2001)
83:217–233.
[132] Reubel G.H., Pekin J., Webb-Wagg K.,
Hardy C.M., Nucleotide sequence of

372 C. Buonavoglia, V. Martella
glycoprotein genes B, C, D, G, H and I,
the thymidine kinase and protein kinase
genes and gene homologue UL24 of an
Australian isolate of canine herpesvirus,
Virus Genes (2002) 25:195–200.
[133] Rijsewijk F.A., Luiten E.J., Daus F.J.,
van der Heijden R.W., van Oirschot J.T.,
Prevalence of antibodies against canine herpesvirus
1 in dogs in The Netherlands in
1997-1998, Vet. Microbiol. (1999) 65:1–7.
[134] Robinson A.J., Crerar S.K., Waight Sharma
N., Muller W.J., Bradley M.P., Prevalence
of serum antibodies to canine adenovirus
and canine herpesvirus in the European red
fox (Vulpes vulpes) in Australia, Aust. Vet.
J. (2005) 83:356–361.
[135] Ronsse V., Verstegen J., Onclin K., Guiot
A.L., Aeberle C., Nauwynck H.J. et al.,
Seroprevalence of canine herpesvirus-1
in the Belgian dog population in 2000,
Reprod. Domest. Anim. (2002) 37:299–
304.
[136] Rosen L., Serologic groupings of reovirus
by hemagglutination inhibition, Am. J.
Hyg. (1960) 71:242–249.
[137] Rosenberg F.J., Lief F.S., Todd J.D., Reif
J.F., Studies on canine respiratory viruses.
I. Experimental infection of dogs with an
SV5-like canine parainfluenza agent, Am.
J. Epidemiol. (1971) 94:147–165.
[138] Rota P.A., Maes R.K., Homology between
feline herpesvirus-1 and canine herpesvirus,
Arch. Virol. (1990) 115:139–145.
[139] Rota P.A., Oberste M.S., Monroe S.S., et
al., Characterization of a novel coronavirus
associated with severe acute respiratory
syndrome, Science (2003) 300:1394–1399.
[140] Rubarth S., An acute virus disease with
liver lesion in dogs (hepatitis contagiosa canis),
Acta Pathol. Microbiol. Scand. (1947)
Suppl. 69:1–207.
[141] Sabin A.B., Reoviruses, Science (1959)
130:1397–1389.
[142] Sagazio P., Cirone F., Pratelli A., Tempesta
M., Buonavoglia D., Sasanelli M., Rubino
G., Infezione da herpesvirus del cane: diffusione
sierologica in Puglia, Obiettivi e
Documenti Veterinari (1998) 5:63–67.
[143] Sanderson C.M., McQueen N.L., Nayak
D.P., Sendai virus assembly: M protein
binds to viral glycoproteins in transit
through the secretory pathway, J. Virol.
(1993) 67:651–663.
[144] Saona-Black L., Lee K.M., Infection
of dogs and cats with canine parainfluenza
virus and the application of a
conglutinating-complement-absorption test
on cat serums, Cornell Vet. (1970) 60:120–
134.
[145] Scott F.W., Kahn D.E., Gillespie J.H.,
Feline reovirus: isolation, characterization,
and pathogenicity of a feline reovirus, Am.
J. Vet. Res. (1970) 31:11–20.
[146] Sen G.C., Viruses and interferons, Annu.
Rev. Microbiol. (2001) 55:255–281.
[147] Skilling D.E., Evermann J.F., Stott J.L.,
Saliki J.T., Trites A.W., Infectious disease
and the decline of Steller sea lions
(Eumetopias jubatus) in Alaska, USA: insights
from serologic data, J. Wildl. Dis.
(2005) 41:512–524.
[148] Southern J.A., Young D.F., Heaney
F., Baumgärtner W.K., Randall R.E.,
Identification of an epitope on the P and V
proteins of simian virus 5 that distinguishes
between two isolates with different biological
characteristics, J. Gen. Virol. (1991)
72:1551–1557.
[149] Sovinova O., Tumova B., Pouska F., Nemec
J., Isolation of a virus causing respiratory
disease in horses, Acta Virol. (1958) 2:52–
61.
[150] Stephensen C.B., Casebolt D.B.,
Gangopadhyay N.N., Phylogenetic analysis
of a highly conserved region of the polymerase
gene from 11 coronaviruses and
development of a consensus polymerase
chain reaction assay, Virus Res. (1999)
60:181–189.
[151] Subbarao K., Klimov A., Katz J., Regnery
H., Lim W., Hall H., Perdue M., Swayne D.,
Bender C., Huang J., Hemphill M., Rowe
T., Shaw M., Xu X., Fukuda K., Cox N.,
Characterization of an avian influenza A
(H5N1) virus isolated from a child with
a fatal respiratory illness, Science (1998)
279:93–96.
[152] Swango L.J., Eddy G.A., Binn L.N.,
Serologic comparisons of infectious canine
hepatitis and Toronto A26/61 canine adenoviruses,
Am. J. Vet. Res. (1969) 30:1381-
1387.
[153] Swango L.J., Wooding W.L., Binn L.N., A
comparison of the pathogenesis and antigenicity
of infectious canine hepatitis virus
and the A26/61 virus strain (Toronto), J.
Am. Vet. Med. Assoc. (1970) 156:1687–
1696.
[154] Tham K.M., Horner G.W., Hunter R.,
Isolation and identification of canine adenovirus
type-2 from the upper respiratory
tract of a dog, N. Z. Vet. J. (1998) 46:102–
105.

[155] Tong S., Li M., Vincent A., Compans R.W.,
Fritsch E., Beier R., Klenk C., Ohuchi M.,
Klenk H.-D., Regulation of fusion activity
by the cytoplasmic domain of a paramyxovirus
F protein, Virology (2002) 301:322–
333.
[156] Truyen U., Muller T., Heidrich R.,
Tackmann K., Carmichael L.E., Survey on
viral pathogens in wild red foxes (Vulpes
vulpes) in Germany with emphasis on parvoviruses
and analysis of a DNA sequence
from a red fox parvovirus, Epidemiol.
Infect. (1998) 121:433-440.
[157] Tyler K.L., Mammalian reoviruses, in:
Knipe D.M., Howley P.M. (Eds.), Fields
Virology, 4th edition, Lippincott Williams
& Wilkins, Philadelphia, 2001, pp. 1729–
1745.
[158] Ueland K., Serological, bacteriological
and clinical observations on an outbreak
of canine infectious tracheobronchitis in
Norway, Vet. Rec. (1990) 126:481–483.
[159] Van Regenmortel M.H.V., Fauquet C.M.,
Bishop D.H.L., Carstens E.B., Estes M.K.,
Lemon S.M., Maniloff J., Mayo M.A.,
McGeoch D.J., Pringle C.R., Wickner
B.R.B., Virus Taxonomy, Seventh Report of
the International Committee on Taxonomy
of Viruses, Academic Press, New York, NY,
2000.
[160] Vennema H., Poland A., Foley J., Pedersen
N.C., Feline infectious peritonitis viruses
arise by mutation from endemic feline
enteric coronaviruses, Virology (1998)
243:150–157.
[161] Vijgen L., Keyaerts E., Moes E., Thoelen
I., Wollants E., Lemey P., Vandamme A.M.,
Van Ranst M., Complete genomic sequence
of human coronavirus OC43: molecular
clock analysis suggests a relatively recent
zoonotic coronavirus transmission event, J.
Virol. 79 (2005) 1595–1604.
[162] Waddell G.H., Teigland M.B., Sigel M.M.,
A new influenza virus associated with
equine respiratory disease, J. Am. Vet.
Med. Assoc. (1963) 143:587–590
[163] Webster R.G., Are equine 1 influenza
viruses still present in horses? Equine Vet.
J. (1993) 25:537–538.
Canine respiratory viruses 373
[164] Webster R.G., The importance of animal influenza
for human disease, Vaccine (2002)
20:16–20.
[165] Weiner H.L., Fields B.N., Neutralization
of reovirus: the gene responsible for the
neutralization antigen, J. Exp. Med. (1977)
146:1305–1310.
[166] Weiner H.L., Raming R.F., Mustoe T.A.,
Fields B.N., Identification of the gene
coding for the hemagglutinin of reovirus,
Virology (1978) 86:581–584.
[167] Weiner H.L., Ault K.A., Fields B.N.,
Interaction of reovirus with cell surface
receptors. I. Murine and human lymphocytes
have a receptor for the emagglutinin
of reovirus type 3, J. Immunol. (1980)
124:2143–2148.
[168] Wigand R., Bartha A., Dreizin R.S.,
Esche H., Ginsberg H.S., Green M.,
Hierholzer J.C., Kalter S.S., McFerran J.B.,
Pettersson U., Russel W.C., Wadell G.,
Adenoviridae: second report, Intervirology
(1982) 18:169–176.
[169] Willoughby K., Bennett M., McCracken
C.M., Gaskell R.M., Molecular phylogenetic
analysis of felid herpesvirus 1, Vet.
Microbiol. (1999) 69:93–97.
[170] Wright P.F., Webster R.G., Orthomyxoviruses,
in: Knipe D.M., Howley P.M.
(Eds.), Fields virology, Lippincott Williams
& Wilkins, Philadelphia, 2001, pp. 1533–
1579.
[171] Yamamoto T., Some physical and growth
characteristics of a canine adenovirus isolated
from dogs with laryngotracheitis,
Can. J. Microbiol. (1966) 12:303–311.
[172] Yamamoto R., Marusyk R.G., Morphological
studies of a canine adenovirus, J.
Gen. Virol. (1968) 2:191–194.
[173] Yoon K.-J., Cooper V.L., Schwartz
K.J., Harmon K.M., Kim W.I., Janke
B.H., Strohbehn J., Butts D., Troutman
J., Influenza virus infection in racing
greyhounds, Emerg. Infect. Dis. (2005)
11:1974–1975.

N o b i v a c ® KC
KC – bredare och snabbare skydd
N o b i v a c ® KC
KC – bredare och snabbare skydd
Nobivac ® KC – produktprofil
N o b i v a c ® K C
• Försvagat levande vaccin för intranasalt
bruk:
• >108.3 cfu B. bronchiseptica stam B-C2
• 103.8 TCID50 CPi
• Dos: 0.4 ml
• Ges i ena näsborren
• Immunitet efter 72 tim

Intranasala vacciner – fördelar
• Inducerar lokal och systemisk
immunitet
• Snabb immunitetsstart
• Fullt skydd efter en enda vaccination
• Kan ges till MDA-positiva valpar*
* Jacobs et al. WSAVA 2006
Lumen Bb
Vaccin Bb
Celler
Plasmacell
Systemisk Immunitet - IgG
IgG
N o b i v a c ® K C
N o b i v a c ® K C
Efter injektions- / intranasal
vaccination:
Vid smitta - Inhalerade Bb
• Fäster till cilierna
• Orsakar infektion
• Inflammationen frisätter IgG
till ytan och cilierna
• Infektion begränsas men
förhindras inte
Lokal immunitet - IgA
IgA
N o b i v a c ® K C
Bb Intranasalt vaccin
administreras på mukosan
som vid naturlig infektion:
• Submukösa plasmaceller
stimuleras till IgA-produktion
• IgA frisätts i mukosan
• Vid smitta - IgA binder
inhalerade Bordetella
bronchiseptica

N o b i v a c ® K C
Intranasala vacciner – nackdelar
• Ibland svårt att administrera
• Kan förekomma nysningar efter
vaccination
• Vaccinerade hundar kan utskilja Bb
(apatogen)
Effektivietsstudie
1 års immunitet
• 12 hundar vacc vid 3 v ålder
• + 6 kontroller samma ålder
N o b i v a c ® K C
• Challenge med Bb och CPi efter 56 veckor
• + 6 10v valpkontroller
Jacobs et al. 2005 Vet Rec, 157, 19-23.
• Provtagna och observerade i 3 veckor
Effektivietsstudie
1 års immunitet – resultat
Efter vacc - signifikant lägre:
- virusisolering (p=0,05)
- mv kliniska poäng (61%, p=0,009)
N o b i v a c ® K C
- mv kliniska poäng valpar (90%, p=0,001)
Jacobs et al. 2005 Vet Rec, 157, 19-23.

Effektivitetsstudie
skydd på 72 tim
• Valpar vacc. vid 8 v
– 20 vacc. (10 + 10) + 10 kontroller
• Challenge med Bb 108,4 CFU/ml i
2ml aerosol efter 48 eller 72 tim
• Bedömning av kliniska symptom
Effektivitetsstudie
skydd på 72 tim – resultat
25
20
15
10
5
0
4
N o b i v a c ® K C
Gore et al. (2005) Vet Rec 156, 482-483
Kliniska poäng efter Bb Challenge - mv
18
Vaccinates (n=10) Controls (n=10)
N o b i v a c ® K C
Kliniska symptom minskade med 73%, signifikant (p=0.002)
Excellent säkerhetsprofil
• Experimentella säkerhetsstudier:
2-veckors valpar x 1
2-veckors valpar x 2
2-veckors valpar x 10
Dräktiga i alla tre trimestrar
• Resultat:
Inget onormalt!
N o b i v a c ® K C

Excellent säkerhetsprofil
N o b i v a c ® K C
• Fältstudier:
- 1289 hundar varierande ålder, ras och kön
(varav 104 st dräktiga)
- observerades i 2 veckor efter vaccinering
• Resultat:
0,1 % visade
milda, övergående
symptom
N o b i v a c ® K C
Nobivac ® KC – sammanfattning
• Bivalent skydd mot kennelhosta
• Bra skydd vid Bordetella-challenge
• Sgs ingen virusutsöndring efter CPi-challenge
• Endast EN dos
• Mycket snabb immunitetsstart Bb – 72h!
• Duration minst 12 mån
• Kan användas ihop med andra Nobivac-vacc*
• Excellent säkerhetsprofil – 2v valpar, dräktiga
* Jacobs et al. Vet Rec (2007) Jan 13, 160, 41-45.
N o b i v a c ® K C
Vilka hundar ska vaccineras mot
kennelhosta?
• Både SVS/SVAs och WSAVAs
expertgrupper rek. vaccination till hundar
som vistas i hundrika miljöer
• SKK rek. Vaccination inför utställningar,
prov och tävlingar

N o b i v a c ® K C
Nobivac ® KC – vaccinationsteknik
• Låt ägaren stå framför hunden
• Stå själv bakom hunden
• Håll om nosen med ena handen
• Droppa i dropparna, inte spraya
Förpackningar och priser
Apotek IntervetDirekt
5 doser 310:- -
25 doser 1385:- 1299:-
N o b i v a c ® K C
N o b i v a c ® K C
KC – bredare och snabbare skydd
Anv Använd Anv nd Nobivac ® KC till
hundar i alla åldrar ldrar när ett
bredare och snabbare skydd
mot kennelhosta önskas nskas nskas!

Svenska erfarenheter av vaccination med intranasalt kennelhostevaccin
Anette Johansson, distriktsveterinär i Kiruna
Det är femtonde året i rad som vi vaccinerar våra hundar samt en del andra slädhundar med
intranasalt kennelhostevaccin.
Första gången var 1994 och då fick hundarna två vaccinationer med en månads intervall. Efter
varje vaccination fick de vila en vecka. Under de första åren vaccinerade vi två gånger. De
fick en första vaccination tidigt på hösten och en andra i december i god tid innan
tävlingssäsongen drog igång på allvar.
Under de följande tio åren har de vaccinerats bara en gång per säsong och det har gett ett
tillräckligt skydd. Vi vaccinerar under perioden oktober till december. Eftersom hundarna får
vila några dagar (oftast 5 dagar) har vi vaccinerat när det har blivit isigt före och omöjligt att
träna eller om vi har varit bortresta.
Från början var det ett vaccin av annat fabrikat men nu har vi använt Nobivac KC i flera år.
Det intranasala vaccinet ger ett mycket bra skydd. Sedan vi började använda intranasalt
vaccin har våra vaccinerade hundar inte drabbats av kennelhosta trots att smittan har grasserat
i omgivningarna. Vi har oftast inte vaccinerat valpar och så sent som i vintras smittades
valparna när grannarnas hundar hade hosta. Men ingen av de vuxna hundarna fick hosta.
Risken för biverkningar lär vara högre med intranasalt vaccin och det är därför vi bara har
vaccinerat de vuxna hundarna. Det är också anledningen till att hundarna får vila från
träningen veckan efter vaccination. Det är nog också viktigt att hundarna är i god kondition
och friska för övrigt.
Genom åren har någon enstaka hund hostat efter vaccinationen. Det har varit hundar som inte
tidigare har varit vaccinerade med intranasalt vaccin. Det är enda biverkningarna vi har sett på
våra hundar.
Däremot har jag ett par fall av allvarligare biverkningar på två andra kennlar.
Det första var ett spann som skulle tävla på medeldistanstävlingen Alpirod som gick i
Alperna. Det var redan 1992 och första gången jag sökte licens på intranasalt vaccin.
Hundarna vaccinerades en tid innan de skulle åka på tävlingen. De blev riktigt sjuka med
hosta och andra symtom från luftvägarna. En del fick pneumoni. Förmodligen var inte
hundarna i tillräckligt god kondition eller kanske rentav nerkörda av för mycket träning. Det
var ingen mönsterkennel vad gäller utfodring och kennelmiljö.
En annan kennel med ca 20 hundar vaccinerades i januari månad för ca 10 år sedan.
Hundhållningen var bra och träningen seriös. Ändå blev flertalet av hundarna sjuka med hosta
och luftvägssymtom. Även då fick någon hund lunginflammation. Kennelhosta grasserade
redan när hundarna vaccinerades. Sannolikt var de redan smittade och vaccinationen gjorde
utbrottet allvarligare.
Jag har mycket goda erfarenheter av det intranasala vaccinet. Jag vet inte om risken för
biverkningar är mindre idag än tidigare men jag rekommenderar att man vaccinerar i god tid
innan säsongen börjar på allvar, att hundarna är i god kondition och att man erbjuder dem en
viloperiod efter vaccinationen.

Jämförande studie mellan vaccinationsantikroppar mot rabies.
Louise Treiberg Berndtsson, leg.vet, enheten för virologi, immunbiologi och parasitologi, SVA.
Bakgrund.
I och med Sveriges inträde i EU fick hundar och katter möjlighet att utan karantän föras in i Sverige.
Djuren måste dock vara vaccinerade mot rabies och dessutom kontrollerade att de erhållit en skyddande titer
på minst 0,5 IE/ml serum 120 dagar efter senaste vaccination.
SVA har sedan detta blev möjligt tagit emot blodprov/serum för analys. Analysen görs i enlighet med OIEs
och EUs community reference laboratory, CRL instruktioner, med deltagande i årliga ringtester.
SVA testar årligen flera tusen prover för rabies antikroppar. De första åren fanns i Sverige bara ett godkänt
vaccin, Rabisin Vet, men sedan 1998 finns 2 st godkända vaccin då även Nobivac Rabies Vet är inregistrerat i
Sverige.
Vi tyckte oss märka efter en tid att det var fler hundar som inte blev godkända om de var vaccinerade med
Nobivac Rabies Vet än de som var vaccinerade med Rabisin vet. Vi fick också indikationer från veterinär och
kliniker att fler hundar än tidigare inte blivit godkända.
Vi visste sedan tidigare att det skilde sig om djuret var vaccinerat en eller två gånger samt om vaccinet även
innehöll komponenten Leptospiros.
Med hjälp av medel från Intervet och Merial beslöts att en undersökning av prov tagna under ett år skulle
undersökas huruvida det var någon skillnad mellan vaccinerna. Vi beslöt också att samtidigt undersöka om
ålder, kön, ras samt om djuret är vaccinerat en eller två gånger har betydelse.
Material
Serum från 6.881 hundar, provtagna i Sverige analyserade år 2005 på SVA, avdelningen för virologi. Prov har
skickats in från hela landet av olika veterinärer och kliniker. Av de 6.881 hundarna som ingår i studien var
3.584 st (52,08%) vaccinerade med Nobivac Rabies Vet och 3.297 st (47,91%) vaccinerade med Rabisin vet.
Analysmetod
Serumproverna analyserades med FAVN-testen, en modifierad serumneutralisations test utarbetad av
AFSSA, Nancy, OIEs referenslab och EUs CRL-lab. (Cliquet et al, 1998).
Resultat
Vid studien kunde några faktorer påvisas som har stor betydelse för om hunden ska klara gränsen på
0,5IE/ml eller ej.
1. En eller två immuniseringar.
Dubbla vaccineringar har stor betydelse för om hunden ska klara gränsen eller ej om hunden är
vaccinerad med Nobivac ® Rabies Vet.
2. Hundens ålder
Hundar under ett års ålder klarade sig något sämre än äldre hundar om de vaccineras med Nobivac
Rabies Vet.
3. Skillnader mellan de 2 olika vaccinerna
Av de 3.584 st hundarna vaccinerade med Nobivac Rabies Vet klarade 87,28% gränsen 0,5 IE/ml
Av de 3.297 st hundarna vaccinerade med Rabisin Vet klarade 97,15% gränsen 0,5 IE/ml
Kön på hunden spelade ingen roll.

Varför skillnad i resultat?
Vaccinerna är framställda av olika virusstammar. Nobivac ® Rabies vet innehåller RIV från Pasteur Institutet
och Rabisin® Vet innehåller Wistar G 57 också från Pasteur Institutet. I analysmetoden används CVS-11
virus som kan vara närmare besläktat med Wistar G 57 (likheter i G-proteinet) och därmed lättare fångar upp
antikroppar riktade mot detta virus.

Abstract
The influence of homologous vs. heterologous challenge
virus strains on the serological test results of rabies
virus neutralizing assays
Susan M. Moore*, Teri A. Ricke, Rolan D. Davis, Deborah J. Briggs
Department of Diagnostic Medicine, College of Veterinary Medicine, Kansas State University, Manhattan, KS 66506, USA
Received 19 May 2005; accepted 12 June 2005
The effect that the relatedness of the viral seed strain used to produce rabies vaccines has to the strain of challenge virus used to
measure rabies virus neutralizing antibodies after vaccination was evaluated. Serum samples from 173 subjects vaccinated with
either purified Vero cell rabies vaccine (PVRV), produced from the Pittman Moore (PM) seed strain of rabies virus, or purified chick
embryo cell rabies vaccine (PCECV), produced from the Flury low egg passage (Flury-LEP) seed strain of rabies virus, were tested
in parallel assays by RFFIT using a homologous and a heterologous testing system. In the homologous system, CVS-11 was used as
the challenge virus in the assay to evaluate the humoral immune response in subjects vaccinated with PVRV and Flury-LEP was
used for subjects vaccinated with PCECV. In the heterologous system, CVS-11 was used as the challenge virus in the assay to
evaluate subjects vaccinated with PCECV and Flury-LEP was used for subjects vaccinated with PVRV. Although the difference in G
protein homology between the CVS-11 and Flury-LEP rabies virus strains has been reported to be only 5.8%, the use of
a homologous testing system resulted in approximately 30% higher titers for nearly two-thirds of the samples from both vaccine
groups compared to a heterologous testing system. The evaluation of equivalence of the immune response after vaccination with the
two different vaccines was dependent upon the type of testing system, homologous or heterologous, used to evaluate the level of
rabies virus neutralizing antibodies. Equivalence between the vaccines was achieved when a homologous testing system was used but
not when a heterologous testing system was used. The results of this study indicate that the strain of virus used in the biological
assays to measure the level of rabies virus neutralizing antibodies after vaccination could profoundly influence the evaluation of
rabies vaccines.
Ó 2005 The International Association for Biologicals. Published by Elsevier Ltd. All rights reserved.
Keywords: CVS-11; Flury-LEP; RFFIT; Serological testing; Rabies vaccine
1. Introduction
Biologicals 33 (2005) 269e276
The immune response to rabies vaccination involves
activation of rabies virus-specific B cells which differentiate
into plasma cells producing antibody and memory
B cells. Although antibodies specific for the rabies virus
glycoprotein (G) and nucleoprotein (N) (as well as other
* Corresponding author. Tel.: C1 785 532 5650; fax: C1 785 532 4474.
E-mail address: smoore@vet.ksu.edu (S.M. Moore).
rabies viral proteins) are produced after vaccination,
published reports indicate that it is the antibodies
specifically directed against antigenic components of
the G protein that neutralize the rabies virus [1]. Rabies
virus-specific CD4C T cells, primarily induced by the
rabies virus N protein, assist in B cell immunoglobulin
class switching and immunoglobulin production. Due to
the lack of a well-established practical method to
measure the cellular immune response against rabies
virus and because rabies virus neutralizing antibodies
1045-1056/05/$30.00 Ó 2005 The International Association for Biologicals. Published by Elsevier Ltd. All rights reserved.
doi:10.1016/j.biologicals.2005.06.005
www.elsevier.com/locate/biologicals

270 S.M. Moore et al. / Biologicals 33 (2005) 269e276
(RVNA) are critical for protection against rabies
infection, the standard method to verify that an immune
response has occurred after rabies vaccination is to
evaluate the level of RVNA in sera. The World Health
Organization (WHO) recognizes two RVNA assays to
assess the humoral immune response after rabies
vaccination: the Rapid Fluorescent Focus Inhibition
Test (RFFIT) and the Fluorescent Antibody Virus
Neutralization Test (FAVN). Both assays utilize the
Challenge Virus Standard (CVS-11) strain of rabies
virus as the challenge virus to quantitate the neutralization
activity of RVNA produced in response to
a rabies vaccine [2,3]. Previous studies have demonstrated
the significant influence that the strain of
challenge virus used in an assay has on the measurement
of vaccine potency [4,5]. Indeed, published reports
indicate that higher vaccine potency values are achieved
when a homologous challenge virus is used for potency
testing as compared to when a heterologous challenge
virus strain is used. A similar effect has been demonstrated
in the serological test results from serum samples
assayed for the presence of specific antibody against
different genotypes of lyssaviruses including rabies virus.
For example, higher RVNA titer values were obtained
against rabies virus, (lyssavirus genotype 1) as opposed
to lyssavirus genotypes 2e7 when the source of the
antibody that was evaluated was pooled sera from
persons vaccinated against rabies virus (lyssavirus
genotype 1) [6]. Additionally, another study reported
variations in RVNA titer values when two different CVS
strains were used as the challenge virus [7].
There are several cell culture rabies vaccines licensed
for use throughout the world. Many of these vaccines
are produced from different rabies virus seed strains
including: Pittman Moore (PM) rabies virus strain used
to produce human diploid cell rabies vaccine (HDCV),
purified Vero cell rabies vaccine (PVRV) and purified
duck embryo cell rabies vaccine (PDEV); Flury high egg
passage (Flury-HEP) or Flury low egg passage (Flury-
LEP) rabies virus strain used to produce two different
types of purified chick embryo cell rabies vaccine
(PCECV); and Kissling rabies virus strain of Challenge
Virus Standard used to produce rabies vaccine adsorbed
(RVA). The PM and Kissling rabies virus strains
originated from the brain of a rabid cow in France in
1882 and the Flury-LEP strain originated from a human
patient in the USA who died of rabies in 1939.
Investigation of the phylogenetic trees of the G and N
rabies virus proteins originating from different vaccine
seed strains indicates a much closer relationship exists
between the PM and CVS strains of rabies virus than
between the Flury-LEP and the CVS strains (Fig. 1).
Published reports also indicate areas of differences exist
between the amino acid sequence of the G protein of
CVS and PM and the G protein of CVS and Flury-LEP
rabies virus strains (Fig. 2). It is important to note that
there are no amino acid sequence differences in the
known, mapped antigenic sites [8]. However, six of the
eight known antigenic sites (epitopes) of the G protein
are conformational and any amino acid changes in close
proximity to these epitopes could potentially affect the
folding of the protein [8]. Additionally, the transmembrane
region has been reported to affect folding of the
ectodomain resulting in subtle conformational changes
of the antigenic sites [9]. The production of RVNA
involves a process of fine-tuning of specificity resulting in
the selection of B cell clones with the highest avidity to
a specific antigen. The potential differences in the
G protein antigenic sites of the original seed virus strains
used in the production of the different rabies vaccines
could result in the preferential production of antibodies
with the highest affinity for antigenic sites resembling the
vaccine seed virus strain. Thus, the strain of challenge
virus used in an RVNA assay and the type of vaccine that
a person was vaccinated with could profoundly influence
the serological test results after vaccination. If this is
correct, RVNA assays using homologous testing systems
(wherein the strain of challenge virus used in the testing
assay is very closely related to the seed virus strain used
to produce the vaccine that a subject received) would
report higher titer values than heterologous testing
systems (wherein the strain of challenge virus used in
the testing system is less closely related to the seed virus
strain used to produce the vaccine that a subject
received). The following study was conducted to determine
the influence that the strain of rabies virus
(homologous vs. heterologous) used as the challenge
virus in a serological assay and the strain of seed virus
used in the production of the rabies vaccine that a subject
received has on the quantitative evaluation of RVNA.
2. Materials and methods
2.1. Challenge virus
Two strains of rabies virus were evaluated as the
challenge virus in the RFFIT assays used to quantitate
the amount of RVNA present in serum samples. The
CVS-11 strain was obtained from the Centers for
Disease Control and Prevention (Atlanta, GA). Seed
virus of the CVS-11 was grown on BHK cells to produce
stock virus. The Flury-LEP strain was obtained from
Chiron Vaccines (Marburg, Germany); stock virus was
grown in primary chicken fibroblasts. Stock virus
preparations were titered to obtain a working dilution
of 50 TCID50.
2.2. Serum samples
Serum samples used in the analyses were obtained
from subjects who had received the same simulated

post-exposure vaccination regimen with either PCECV
(n Z 86) or PVRV (n Z 87). Subjects did not receive
rabies immune globulin (RIG). Serum samples that were
collected on day 14 and day 90 after initial vaccination
were included in the study. Serum samples were
randomly placed into five testing groups (Groups 1
through 5). Each group contained from 60 to 120 serum
samples including samples from subjects vaccinated with
PCECV as well as subjects vaccinated with PVRV. All
serum samples were coded to ensure that testing was
conducted blindly and unsorted by vaccine group.
2.3. Equilibration of working dilution
of challenge virus
The working dilution of the challenge virus was
equilibrated to 50 TCID50 for both the CVS-11 and the
Flury-LEP rabies virus strains. The titer of the challenge
virus was calculated for each test set of serological
samples in order to assure equivalence in testing criteria.
For all test runs, the titer of the challenge virus was
maintained within one standard deviation of the average
S.M. Moore et al. / Biologicals 33 (2005) 269e276
Fig. 1. Phylogenetic relationship of rabies virus strains (courtesy of Dr Iris Stalkamp, Institut fu¨r Virologie, Giessen, Germany).
calculation (41.1 TCID50) for virus titer throughout the
entire evaluation.
2.4. Serological testing
The RFFIT, using CVS-11 and Flury-LEP as the
challenge virus strains in parallel, was used to assay all
serum samples, as previously described [10]. Briefly,
100 mL of each serum sample, in duplicate, was diluted
in serial fivefold dilutions and loaded into 8-well Labtek
chamber slides after which 100 mL of the challenge
virus, at a concentration of 50 TCID50, was added.
Slides were incubated at 37 C for 90 min after which
200 mL of a suspension of 5 ! 10 5 BHK cells was added
to each well. Slides were placed in a 5% CO2 incubator
at 37 C for 24 h. After incubation the slides were
washed and fixed in 80% cold acetone, dried and stained
with FITC conjugated anti-rabies antibody (Chemicon,
Temecula, CA). Twenty fields/well were examined under
160! magnification using a fluorescence microscope for
the presence of rabies virus and RVNA titers were
calculated using the Reed and Muench method. Reciprocal
titers were used in the evaluations in order to
271

272 S.M. Moore et al. / Biologicals 33 (2005) 269e276
Fig. 2. Amino acid alignment of the rabies glycoprotein from Flury, CVS and PM strains (courtesy of Dr Iris Stalkamp, Institut fu¨r Virologie,
Giessen, Germany). There are fewer amino acid sequence changes from CVS to PM (filled arrows) than CVS to Flury (open arrows). The changes are
not in areas of the mapped antigenic sites of the rabies glycoprotein (shaded triangles). The transmembrane sequence is indicated by the boxed area.
eliminate the need to calculate international units using
titer results obtained from an international rabies
reference serum that originated from subjects only
vaccinated with a rabies vaccine produced from a PM
seed strain of rabies virus.
2.5. Statistical analyses
After all serum samples were tested separately with
both the CVS-11 and the Flury-LEP rabies challenge
virus strains, the identification of the two vaccination
groups (PVRV and PCECV) was unblinded and the
RVNA titers were statistically analyzed to determine the
effect of serological testing by means of a homologous
vs. heterologous test. To determine whether any straindependent
difference in neutralizing antibody was
magnified at higher titers, the titer results (both day 14
and day 90) were sorted into response groups, the
geometric mean titer (GMT) of the groups was
calculated, and the GMT by challenge virus was
compared. Additionally, to determine whether maturation
of the antibody response amplified the differences in
GMT, the titer responses by day of serum drawn were
sorted and the GMT of the groups was calculated and
compared by challenge virus.
3. Results
The virus titer of CVS-11 and Flury-LEP, used as the
challenge virus in each of the five serological testing
groups, remained consistently equivalent throughout the
testing period (Table 1). There was a similar wide range
of RVNA titers obtained for each vaccination group,
independent of whether CVS-11 or Flury-LEP was used
as the challenge virus strain (Table 2). There were two
outlier reciprocal titer values in the PVRV vaccination
group, 9500 and 19 700, exhibited by the same subject

Table 1
Titer of Challenge Virus Standard (CVS-11) and Flury low egg passage
(Flury-LEP), the two rabies virus strains used as the challenge viruses
for each serological testing group
Serological testing group Titer of CVS-11 Titer of Flury-LEP
1 42.1 41.1
2 40.0 41.2
3 41.0 40.0
4 41.4 42.3
5 41.1 40.9
Geometric mean 41.1 41.1
Virus titer is expressed in TCID50. on days 14 and 90, respectively. The GMT for each
group indicates higher RVNA titers were reported when
a homologous challenge virus strain was used in the
serological assay. The RVNA test results of individual
serum samples indicated that there was a clear trend to
report higher titers when a homologous testing system
(CVS-11 used as the challenge virus in the RFFIT for
testing sera from subjects vaccinated with PVRV and
Flury-LEP used as the challenge virus in the RFFIT for
testing sera from subjects vaccinated with PCECV)
rather than a heterologous testing system (Flury-LEP
used as the challenge virus in the RFFIT for testing sera
from subjects vaccinated with PVRV and CVS-11 used
as the challenge virus in the RFFIT for testing sera from
subjects vaccinated with PCECV) was used (Fig. 3).
The RVNA values in both the PCECV and the
PVRV vaccine groups included titers in the low, medium
and high range, regardless of which challenge virus was
used in the assay. Low, medium, and high ranges were
designated for this set of results to determine possible
trends associated with the strength of the antibody
response. Nearly two-thirds of the samples from each
Table 2
Rabies virus neutralizing antibody (RVNA) titers from the Rapid
Fluorescent Focus Inhibition Test (RFFIT) using a homologous
challenge virus testing system and a heterologous challenge virus
testing system
Vaccine RFFIT testing GMT (range)
administered system
Challenge virus Day 14 Day 90
PCECV Homologous 1855 265
Flury-LEP (320e6300) (45e1500)
Heterologous 1275 183
CVS-11 (145e5400) (45e1100)
PVRV Homologous 2364 274
CVS-11 (360e8500) (45e9500)
Heterologous 1448 188
Flury-LEP (70e8500) (45e19700)
Serum samples were obtained from subjects vaccinated with purified
chick embryo cell rabies vaccine (PCECV) or purified Vero cell rabies
vaccine (PVRV) and were assayed using CVS-11 and Flury-LEP as the
challenge viruses in the RFFIT.
S.M. Moore et al. / Biologicals 33 (2005) 269e276
titers (Flury) log dil
4.00
3.50
3.00
2.50
2.00
1.50
1.50 2.00 2.50 3.00 3.50 4.00
vaccine group reported higher titers when a homologous
challenge virus strain was used for the RFFIT assay,
63% for PCECV and 65% for PVRV. Approximately
30% of the serum samples tested in each vaccine group
reported titers that were the same value or similar
(within one standard deviation) regardless of whether
they were assayed using a homologous or a heterologous
challenge virus strain.
The percent reduction of reported RVNA titers,
when switching from a homologous testing system to
a heterologous testing system was 23%, 47%, and 33%,
respectively, for the low, medium, and high response
groups in the PVRV vaccination group and 27%, 25%,
and 40% in the PCECV vaccination group (Fig. 4).
Thus, there was no clear trend of higher or lower RVNA
titers related to the type of testing system, and whether
the serum tested belonged to the low, medium, or high
response group. The overall reduction in RVNA titer
values when switching from a homologous challenge
virus assay to a heterologous challenge virus assay was
33% for the PVRV vaccination group, and a 31%
reduction for the PCECV vaccination group.
On both day 14 (data not shown) and day 90 the
GMTs were higher when a homologous challenge virus
system was used for the PVRV and PCECV vaccination
groups (Fig. 5).
4. Discussion
PCECV Subjects PVRV Subjects
titers (CVS) log dil
Fig. 3. Serum samples from day 90 after administration of purified
chick embryo cell rabies vaccine (PCECV) or purified Vero cell rabies
vaccine (PVRV) given in a post-exposure prophylaxis regimen were
analyzed twice by RFFIT. In one assay CVS-11 was utilized as the
challenge virus in the RFFIT and in the second RFFIT, Flury-LEP
was utilized as the challenge virus. The rabies virus neutralization titer
(RVNA) result obtained for each serum sample was plotted according
to the challenge virus used in the RFFIT. The line of unity represents
expected RFFIT values that would be equivalent regardless of whether
CVS-11 or Flury-LEP rabies virus was used as the challenge virus for
patients vaccinated with PCECV or PVRV. Similar results were seen
with the day 14 results (data not shown).
Neutralizing antibodies play a critical role in immune
protection against rabies infection. Therefore, it is
273

274 S.M. Moore et al. / Biologicals 33 (2005) 269e276
GMT
260
240
220
200
180
160
140
120
100
80
60
PCECV homologous
PCECV heterologous
Low
PVRV homologous
PVRV heterologous
1200
1100
1000
900
800
700
600
appropriate to utilize RVNA assays to measure the
immune response after rabies vaccination rather than
relying on antigen-binding assays, which do not measure
the function of the antibodies produced. Indeed,
currently the most accepted method for measuring the
immune response to rabies antigen is to measure
the amount of RVNA in serum. In the United States,
the Advisory Committee on Immunization Practices
(ACIP) recommends that RVNA testing should be
performed using a virus neutralization assay; those
persons at risk of contracting rabies should have their
RVNA levels measured periodically; and a booster
should be administered to persons at risk of contracting
rabies when their RVNA titer falls below complete
neutralization of a specific quality of rabies virus at a 1:5
500
400
300
200
PCECV homologous
PCECV heterologous
Medium
PVRV homologous
PVRV heterologous
6000
5500
5000
4500
4000
3500
3000
2500
2000
1500
1000
PCECV homologous
PCECV heterologous
High
PVRV homologous
PVRV heterologous
Fig. 4. Depicted are the geometric mean titers (GMT) of serum samples analyzed by the Rapid Fluorescent Focus Inhibition Test (RFFIT) and
separated into high, medium and low titer results. Serum samples were collected from subjects vaccinated with purified Vero cell rabies vaccine
(PVRV) or purified chick embryo cell rabies vaccine (PCECV), and tested with RFFIT in either a homologous or heterologous testing system. A
homologous testing system included sera from subjects vaccinated with PCECV and analyzed by RFFIT using the Flury-LEP as the challenge and
sera from subjects vaccinated with PVRV and analyzed by RFFIT using the CVS-11 as the challenge virus. A heterologous testing system included
sera from subjects vaccinated with PCECV and analyzed by RFFIT using the CVS-11 challenge virus and sera from subjects vaccinated with PVRV
and analyzed by RFFIT using the Flury-LEP as the challenge virus.
GMT (reciprocal titers)
1000
100
10
Challenge Virus Strain:
GMR
(PCECV/PVRV)
PCECV heterologous
PCECV homologous
serum dilution by the RFFIT (the World Health
Organization recognizes this level to be 0.5 IU/mL)
[11,12]. The evaluation of serological levels of RVNA is
also appropriate for patients who may have a questionable
immune response after post-exposure prophylaxis,
i.e. when vaccine was administered or stored inappropriately,
when a patient may be immunosuppressed, or
when a patient may have had a severe adverse reaction
to the vaccine. Finally, new rabies vaccines are
evaluated, licensed and approved for use partly by
assessing the level of RVNA produced after vaccination
in human subjects enrolled in clinical trials.
As mentioned earlier, the CVS-11 strain of rabies virus,
generally used as the challenge virus in the RFFIT assays
that are used to measure RVNA, differs in how closely it is
PVRV heterologous
PVRV homologous
CVS-11 Flury LEP CVS-11 Flury LEP
0.69 (0.58 - 0.84)
1.48 (1.23 - 1.80)
1.00 (0.83 - 1.21)
Fig. 5. Serum samples from day 90 after administration of purified chick embryo cell rabies vaccine (PCECV) or purified Vero cell rabies vaccine
(PVRV) given in a post-exposure prophylaxis regimen were analyzed by the Rapid Fluorescent Focus Inhibition Test (RFFIT) for rabies virus
neutralizing antibodies, using different challenge virus strains. Depicted are geometric means of reciprocal titers (GMT), error bars represent 90%
confidence intervals. Geometric mean ratios (PCECV/PVRV) of the different challenge strain comparison groups were calculated (90% confidence
intervals in parentheses), resulting in equivalent titers when using the homologous challenge strain Flury-LEP for PCECV and CVS-11 for PVRV.

elated to the PM and Flury strains of rabies viruses that
are used in the production of human rabies vaccines
(Figs. 1 and 2). These differences are in some instances
located in areas that are in close enough proximity to the
antigenic sites (and also in the transmembrane region) to
potentially affect the conformation of the antigenic sites.
It is possible that the differences between the strains of
seed virus used in the production of rabies vaccines are
enough to cause slight conformational changes in the
antigen-binding site of the antibodies that are induced
after vaccination. These slight differences in the antigenbinding
site could cause the antibody to have a higher
affinity for a challenge virus used in an in vitro assay
that more closely resembles the antigen that caused its
production in the first place. The results of our study
indicate that the degree of homology between the strain of
challenge virus used in the RFFIT to measure the immune
response after vaccination and the strain of seed
virus used to produce the vaccine that subjects received
profoundly affects the reported RVNA values. Indeed,
the use of challenge virus strains with equivalent titers in
RFFIT assays resulted in approximately 30% higher
RVNA values in two-thirds of the serum samples we
analyzed when a homologous testing system was used. In
addition, the level of the RVNA titer (high, medium or
low) had no obvious or consistent effect on the percentage
of titer difference reported between the testing systems.
In most cases the choice of the challenge virus strain
used in a rabies virus neutralization assay does not play
a critical role in the evaluation of RVNA titers; for
example, periodic titer evaluations and the determination
of an immune response after post-exposure prophylaxis
where the exact titer level is less important
than the actual detection of neutralizing antibody. In
addition, the strain of rabies virus used in a rabies virus
neutralization assay is unlikely to be a determining
factor in the measurement of RVNA titers in persons
whose pre-exposure series may be from one vaccine
source and subsequent booster(s) from another source.
Similarly, persons who have had a rabies exposure will
have an immune response to the rabies antigens in the
exposure strain and to the vaccine strain confounding
the mix of antibodies produced. For all of the above
mentioned reasons it would provide little benefit to
routinely measure the RVNA response using separate
rabies virus challenge strains. In contrast, the measurement
of the humoral immune response after vaccination
for the specific purpose of evaluating a rabies vaccine
makes the choice of the challenge virus used in a rabies
virus neutralizing assay extremely important. When the
RVNA levels produced against vaccines made from two
different parent strains are compared using an assay that
employs a particular challenge virus strain in the testing
system, the combined effect of the quantity, functionality
and specificity of the respective antibody response
is measured. As demonstrated by this study, if the
S.M. Moore et al. / Biologicals 33 (2005) 269e276
challenge virus used in the assay is more closely related
to one parent virus strain than to the other, the titer
results obtained will be biased toward the homologous
vaccine. Most importantly, the evaluation methods used
to confirm an absence of significant difference between
the immune response produced by two vaccines involve
statistical comparisons of the GMT by the geometric
mean ratio (GMR). The Food and Drug Administration
(FDA) defines bio-equivalence as ‘‘pharmaceutical
equivalents whose rate and extent of absorption are
not statistically different when administered to patients
or subjects at the same molar dose under similar
experimental conditions’’ [13]. In comparing the statistical
evaluation of each vaccine, the confidence intervals
(CI) of the GMR are examined. When the lower limit of
the 95% CI is greater than 50% and the interval
includes 100%, ‘‘non-inferiority’’ is achieved. To determine
the stricter standard of ‘‘bio-equivalence’’, 90%
CI of the GMR must lie within 80e125%. If this
equivalence test is applied for the day 90 results in our
study, the GMTs obtained for PCECV are inferior to
PVRV when serum samples from subjects vaccinated
with PCECV are tested in a heterologous testing system
using the CVS-11 strain of challenge virus. Conversely,
the GMTs obtained for PVRV are inferior to PCECV
when serum samples from subjects vaccinated with
PVRV are tested in a heterologous testing system using
the Flury-LEP strain of challenge virus (Fig. 5). However,
when a homologous testing system is used to test
the serum samples for subjects in each vaccination group,
not only are the two vaccines non-inferior, they are
equivalent.
This report ascertains that the choice of challenge
virus strain used in rabies virus neutralization assays to
evaluate the production of RVNA titers after vaccination
should be taken into consideration when the titer
values will be used for the evaluation of new or existing
vaccines. Clearly, if quantifying the immune response to
the vaccine is the objective, then using a homologous
rabies virus strain in the testing would most appropriately
reflect this goal. Finally, it is important to
remember that modern cell culture rabies vaccines are
highly effective and cross-protection between strains has
been demonstrated [14,15]. The use of a heterologous or
homologous testing system to evaluate the level of
RVNA as a measure of complete ‘protection’ against
rabies infection is incorrect. To date, the level of RVNA
required to be ‘protective’ against infection in humans
is not known for an obvious reason: it is unethical to
conduct challenge experiments in humans to determine
the level of RVNA required for protection. On the
other hand, the use of rabies virus neutralizing antibody
testing systems to measure the immune response to
specific rabies antigens and the response to rabies
vaccines should not only be accurate and precise, but
also meaningful.
275

276 S.M. Moore et al. / Biologicals 33 (2005) 269e276
Acknowledgments
The authors would like to thank the Centers for
Disease Control and Prevention and Chiron Vaccines
for donating the challenge viruses used in this study. In
addition, our appreciation is extended to Dr Iris
Stalkamp for providing phylogenetic data and sequence
comparison information and to Dr Tony Stamp and
Mal Hoover for their valuable assistance in preparing
some of the figures used in this paper.
References
[1] Lafon M, Edelman L, Bouvet JP, Lafage M, Martchatre E.
Human monoclonal antibodies specific for the rabies virus
glycoprotein and N protein. J Gen Virol 1990;71:1689e96.
[2] Smith JS, Yager PA, Baer GM. A rapid reproducible test for
determining rabies neutralizing antibody. Bull World Health
Organ 1973;48:535e41.
[3] Cliquet F, Aubert M, Sagné L. Development of a fluorescent
antibody virus neutralization test (FAVN test) for the quantitation
of rabies-neutralising antibody. J Immunol Methods
1998;212:79e87.
[4] Blancou J, Aubert MF, Cain E, Selve M, Thraenhart O,
Bruckner L. Effect of strain differences on the potency testing of
rabies vaccines in mice. J Biol Stand 1989;17(3):259e66.
[5] Ferguson M, Wachmann B, Needy C, Fitzgerald EA. The effect
of strain differences on the assay of rabies virus glycoprotein by
single radial immunodiffusion. J Biol Stand 1987;15:73e7.
[6] Smith JS. Molecular epidemiology. In: Jackson AC, Wunner WH,
editors. Rabies. 1st ed. San Diego, CA, USA: Academic Press;
2002. p. 79e111.
[7] Smith JS. Rabies serology. In: Baer GM, editor. The natural
history of rabies. 2nd ed. Boca Raton, FL, USA: CRC Press;
1991. p. 235e52.
[8] Tordo N. Characteristics and molecular biology of the rabies
virus. In: Meslin FX, Kaplan MM, Koproski H, editors.
Laboratory techniques in rabies. 4th ed. Geneva, Switzerland:
World Health Organization; 1996. p. 28e51.
[9] Maillard A, Gaudin Y. Rabies virus glycoprotein can fold in two
alternative, antigenically distinct conformations depending on
membrane-anchor type. J Gen Virol 2002;83:1465e76.
[10] Smith JS, Yager PA, Baer GM. A rapid fluorescent focus
inhibition test (RFFIT) for determining rabies virus-neutralizing
antibody. In: Meslin FX, Kaplan MM, Koproski H, editors.
Laboratory techniques in rabies. 4th ed. Geneva, Switzerland:
World Health Organization; 1996. p. 181e92.
[11] Centers for Disease Control and Prevention. Human rabies
prevention-United States, 1999: recommendations of the Advisory
Committee on Immunization Practices (ACIP). MMWR
Morb Mortal Wkly Rep 1999;RR-1:1e21.
[12] WHO Expert Committee on Rabies. 8th Report. Geneva,
Switzerland: World Health Organization; 1992 [WHO Technical
Report Series, No. 824].
[13] Code of Federal Regulations Title 21 Subpart A 320.1 Definitions.
Food and Drug Administration, Health and Human
Services.
[14] Briggs DJ. Public health management of humans at risk. In:
Jackson AC, Wunner WH, editors. Rabies. 1st ed. San Diego,
CA, USA: Academic Press; 2002. p. 401e28.
[15] Lodmell DL, Smith JS, Esposito JJ, Ewalt LC. Cross-protection
of mice against a global spectrum of rabies virus variants. J Virol
1995;69:4957e62.

Nobivac ® Tricat Novum
• Felint panleucopenivirus (stam MW-1)
– 3 års duration
• Felint herpesvirus (stam G2620A)
– Årlig vaccination
• Felint calicivirus (F9)
– Årlig vaccination
Continuum ® Feline HCP
• Intervets kattvaccin i USA
• 3 års DOI* indikation på alla komponenter
• Samma som Nobivac ® Tricat
*Gore et al. Vet Ther 2006
Nobivac ® kattvacciner
Levande attenuerat vaccin
För maximal effekt
11/20/2008Nov 2008
Anna-Karin Lieber
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
2
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
3
1

Varför levande vaccin?
• Stimulerar till cellmedierad immunitet
– Nödvändigt för eliminering av infekterade celler
– Ger längre immunitetsduration
• Effektivare vid närvaro av maternala antikroppar
• Snabbare immunitetsinsättande
– ABCD, WSAWA rekommenderar MLV för katthem
• Replikerar i värdcellen, ger bredare antigenuttryck
– Sannolikt skydd mot fler stammar än med inaktiverat vaccin
Korsneutralisationsstudie (UK)
40 slumpmässigt utvalda isolat testades mot
vaccinstammarna F9 och 255
Ref: Porter et al. (2008) Feline Med Surg 10 (1): 32-40
Korsneutralisationsstudie (UK)
Samma isolat användes för att jämföra
vaccinstammarna F9 och 431/G1
Ref. Lin F et al. (2007) Proceedings at the Voorjaarsdagen 40: 188
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
4
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
5
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
6
2

Nobivac ® kattvacciner
flexibla vacciner med dokumenterat skydd
• Nobivac ® Ducat
– Skydd mot herpes och calici
• Nobivac ® Tricat Novum
– Skydd mot herpes, calici och panleucopeni
• Nobivac ® Forcat
– Skydd mot herpes, calici, panleucopeni och klamydia
Vaccinationsrekommendation
8-9 v
12 v
År 1 booster
År 2 booster
År 3 booster
År 4 booster
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
7
Tricat ® Novum (Forcat)
Tricat ® Novum (Forcat)
Tricat ® Novum (Forcat)
Ducat ® (Forcat)
Ducat ® (Forcat)
Tricat ® Novum (Forcat)
Kvällssymposium: Vaccinering av hund och katt 11/20/200825–27
Nov 2008
8
3

Vaccination av hund och katt 2008
Bitr. stats veterinär Ulrika Windahl
SVA, Enheten för djurhälsa och antibiotikafrågor
Bra vacciner är när de används korrekt kostnadseffektiva läkemedel som förebygger
sjukdom och lidande.
• Vaccinering ingår i en god djurhållning av hund och katt
• Syfte med vaccination: förbättra djurskyddet.
• Kan dock inte kompensera en i övrigt undermålig djurhållning
• En hög andel korrekt vaccinerade individer i en population: minskat generellt smittryck, dvs.
bidrar till att skydda även
o individer som inte svarat på vaccinering
o ännu inte har vaccinerats
o vaccineringen misslyckats
o är dåligt skyddade mot sjukdom (t ex valpar, kattungar)
• ”Ett nej till vaccination mot allvarliga sjukdomar är detsamma som ett ja till kommande
sjukdomsutbrott”
Risk: allvarliga infektioner ”glöms bort” tack vare effektiv vaccinering.
Biverkningsriskerna överdrivs
.
• Ex: parvovirus, HCC tros vara utrotad i Sverige
• Grupper arbetar aktivt (internet) för att motverka vaccinering
• Misstroende mot veterinärer, humansjukvård
• Vetenskapligt underlag för rekommendationer
• Rapportera misstänkta fall av biverkningar!
Vaccination av hund och katt 2008
Ulrika Windahl, SVA

Vaccinering skall liksom annan medicinsk behandling vara individbaserad, baseras
på vetenskap och djurägaren skall vara informerad och involverad
• Vaccin skall användas till rätt djur vid rätt tillfälle = endast vid behov
• Onödiga vaccinationer:
o Mot lindriga infektioner
o Som inte har någon positiv effekt mot infektionsrisk och utveckling av sjukdom
o Infektioner det vaccinerade djuret löper mycket liten risk att drabbas av
o Eller där risken för allvarlig biverkning av vaccinet är större än risken för allvarlig
sjukdom hos det ovaccinerade djuret.
Kunskap krävs om
• Förekomsten av ett smittämne i det land/område djuret vistas i
• Risken för klinisk sjukdom kopplad till det aktuella smittämnet.
• Hur djuret hålls
• Resor
• Krävs att djurägaren är delaktig och tillför nödvändig information.
Djurägaren har också behov av information från veterinär i samband med
vaccinering.
• Djuret har normalt inte har ett gott skydd omedelbart efter förstagångsvaccinationen
• Vilka vacciner som inte skyddar (helt) mot infektion och smittspridning
• Vilken effekt vaccinet förväntas ge; varför det är indicerat ge vaccinet
• Djur som vaccinerats intranasalt kan utsöndra vaccinstammen en kortare tid
• Korrekt information om biverkningar
• Normalt med viss reaktion, jämför barn (och häst)
• Tillägg till besöket: skriftlig information
Vaccination av hund och katt 2008
Ulrika Windahl, SVA

Vaccinationsintervall = hur länge ger en vaccination ett immunologiskt
tillfredställande skydd
• Texten i FassVet = en information från vaccintillverkaren.
o De vaccinationsintervallen = de är de företaget dokumenterat effekten av vid registreringen
av vaccinet.
o Företagen kan inte marknadsföra förhållanden som skiljer sig från de godkända
produktresuméerna.
• Andra studier kan visa på en annan (t.ex. längre varaktighet) av skyddet
• …..och annan effekt
• Veterinära Ansvarsnämnden 2003:
” ……VA i sin bedömning av anmälda fall beaktar så väl vetenskap som beprövad
erfarenhet, varför vaccinationsintervall som bygger på sådan grund inte löper risk
att ifrågasättas på grund av avvikelse från rekommenderade intervall i FassVet.”
• OBS: om en enskild veterinärs vaccinationsrutiner väsentligt avviker från tillverkarens
rekommendationer ökar kravet på aktsamhet, och att veterinären håller sin vetenskapliga kunskap
uppdaterad inom detta område. (Norska vaccinrapp -03)
Bristande möjlighet vaccinera kan leda till
- Dödsfall hos människa
- Dödsfall vilda djur
- Djurlidande, dödsfall husdjur
- Svält…
Generell risk I-länder: för få djur vaccineras – men med onödigt många vacciner.
• Basvaccin (core vaccine) ”till alla”
• Tilläggsvaccin (non-core vaccine) ”Till en del, i en del situationer”
• ”Rekommenderas ej -vacciner” = i ett visst land/generellt
• Vaccination enligt lagstiftning
www.sva.se Djurhälsa – hund, eller katt
SVS-SVA:s vaccinationsrapport 2003, arbete pågår med revidering = uppdatering
ABCD-vets.org
AAFP
AAHA
WSAVA
Vaccination av hund och katt 2008
Ulrika Windahl, SVA

Basvaccin - katt
Basvaccin - hund
Kattungar bas/core vaccine
Tilläggs vaccin ‐ katt
Tilläggsvaccin - hund
Vuxna katter bas/core vaccine
Rekommenderas ej
Rekommenderas ej
Vaccination av hund och katt 2008
Ulrika Windahl, SVA

Valpar bas/core vaccine
Vuxna hundar - bas/core vaccine