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OFFICE INTERNATIONAL DES EPIZOOTIES
W o r l d Organisation for Animal Health
DIAGNOSTIC MANUAL FOR
AQUATIC ANIMAL DISEASES
1995
This Manual has been edited by the OIE Fish Diseases Commission
on the basis of comments received from Member Countries,
and approved by the International Committee of the OIE
OIE Diagnostic Manual for Aquatic Animal Diseases
First Edition, 1995
This Manual is published as a companion volume to the OIE
International Aquatic Animal Health Code
ISBN 92-9044-383-9
© Copyright
OFFICE INTERNATIONAL DES EPIZOOTIES, 1995
12 rue de Prony, 75017 Paris, France
Tel: (33.1)44 15 18 88
Fax: (33.1)42 67 09 87
Telex: EPIZOTI 642 285 F
Reproduction or translation is permitted for non-commercial purposes only,
provided reference is made to the source.
Contents
iii
CONTENTS
General
Foreword
v
Introduction
vii
Abbreviations
ix
Definitions
xi
Acknowledgments
xiii
Diseases of fish
Chapter 1
General information
A.
General bases of fish health surveillance/
1
control programmes
1
B.
Sampling procedures
2
C.
Materials and biological products required for the
isolation and identification of fish pathogens
8
Diseases notifiable to the OIE*
Chapter 2
Epizootic haematopoietic necrosis
21
Chapter 3
Infectious haematopoietic necrosis
31
Chapter 4
Oncorhynchus masou virus disease (Salmonid
herpesvirus type 2)
43
Chapter 5
Spring viraemia of carp
53
Chapter 6
Viral haemorrhagic septicaemia
63
Other significant diseases*
Chapter 7
Channel catfish virus disease (Herpesvirus of
Ictaluridae
type 1)
75
Chapter 8
Viral encephalopathy and retinopathy
85
Chapter 9
Infectious pancreatic necrosis
91
Chapter 10
Infectious salmon anaemia
101
Chapter 11
Epizootic ulcerative syndrome
107
Chapter 12
Bacterial kidney disease (Renibacteriosis)
113
Chapter 13
Enteric septicaemia of catfish (Edwardsiellosis)
127
Chapter 14
Piscirickettsiosis
135
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
iv
C o n t e n t s , cntd.
Diseases of bivalve molluscs
Diseases notifiable to the OIE*
Chapter 15
Diagnostic techniques: general information
141
Chapter 16
Bonamiosis
143
Chapter 17
Haplosporidiosis
147
Chapter 18
Marteiliosis
151
Chapter 19
Mikrocytosis
155
Chapter 20
Perkinsosis
157
Chapter 21
Iridoviroses
161
Diseases of crustaceans
Other significant diseases*
Chapter 22
Baculoviral midgut gland necrosis
Chapter 23
Nuclear polyhedrosis baculoviroses (Penaeus
type baculovirus and Baculovirus
165
monodon-
penaei)
169
Chapter 24
Infectious hypodermal and haematopoietic necrosis .
177
Chapter 25
Yellowhead disease
181
Chapter 26
Crayfish plague
185
List of OEE Reference Laboratories for fish, mollusc
a n d crustacean diseases
193
List of organisations w i t h w h i c h the O I E has
cooperation agreements
195
Diseases notifiable to the OIE' are defined in the International Aquatic Animal Health
Code. They are considered to be of socio-economic and/or public health importance
within countries and also significant in the international trade of aquatic animals and
aquatic animal products, and must be reported to the OIE as specified in the Code.
These diseases were previously known as List B diseases'.
'Other significant diseases' are also defined in the International Aquatic Animal Health
Code. They are of current or potential international significance in aquaculture but have
not been included in the list of diseases notifiable to the OIE because of factors related
to their importance or geographic distribution, or current knowledge about them.
Foreword
v
FOREWORD
The Office International
des Epizooties (OIE) is an
intergovernmental
organisation which was established in 1924 in order to promote world animal
health, and its main activities are as follows:
1
To collect and disseminate
to its Member Countries,
(including
emergency
information)
on the occurrence,
treatment of animal diseases
information
course and
2
To provide guidelines and standards for health regulations
the international trade of animals
3
To promote and co-ordinate research on the pathology, treatment and
prevention of animal diseases when international collaboration in such
research is desirable.
applicable
in
Aquatic animals are included in the concept of 'animals' above.
Diagnostic
procedures for some aquatic animal diseases used to be included in the OIE
International Animal Health Code (1986 edition), but it became clear that a
separate Code and Manual specific to aquatic animal health were needed. The
reasons are that the conditions, problems and requirements in this field are
different to those encountered in other animals, and that international trade in
aquatic animals and their products is intensifying
and increasing
in
importance.
The purpose of this Manual is to provide a uniform approach to the diagnosis
of the diseases listed in the OIE International Aquatic Animal Health Code, so
that the requirements for health certification in connection with trade in
aquatic animals and aquatic animal products, can be met. The Manual is
therefore a companion volume to the Code.
Although many publications exist on the diagnosis and control of aquatic
animal diseases, the OIE Diagnostic Manual for Aquatic Animal Diseases will
hopefully be a key document in describing methods that can be applied in
aquatic animal health laboratories all over the world, thus
increasing
efficiency and promoting improvements in aquatic animal health worldwide.
The task of compiling the Manual was assigned to the OIE Fish Diseases
Commission, and all the chapters were circulated to OIE Member Countries for
comments and revision. The Manual will be continually revised and updated as
new information on aquatic animal diseases in general, and new emerging
diseases in particular, becomes available, and it is intended to publish a new
edition approximately every four years.
Dr Jean Blancou
Director General, OIE
Prof. Tore Hdstein
President, Fish Diseases
Commission
1995
Introduction
vii
INTRODUCTION
The clinical signs in fish with the diseases listed in the OIE
International
Aquatic Animal Health Code are not pathognomonic. Moreover, these
infections may take place as subclinical infections of asymptomatic pathogen
carriers.
The only dependable approach for diagnosis of fish diseases therefore lies in the
specific identification of the pathogens using laboratory methods. These
methods, which are suitable for the diagnosis of isolated cases of disease as part
of the operating of national aquatic animal health surveillance/control
programmes, form the main contents of this Manual.
Basically such health surveillance programmes aim to infer, from the results
provided by standardised laboratory procedures performed with samples
collected according to defined rules, the health status of aquatic animal stocks
from a particular production site and even a geographic zone or entire country.
The satisfactory implementation of such aquatic animal health surveillance/
control programmes, requires the existence of both adequate legislation and
resources in each country interested in aquatic animal health.
The diagnostic methods presented in this Manual are all direct diagnostic
methods. Due to insufficient development of the serological methodology, the
detection of fish antibodies to viruses has not thus far been accepted as a routine
diagnostic method for assessing the health status of fish populations. However,
the validation of some serological techniques for diagnosis of certain fish virus
infections could arise in the near future, rendering the use of serology more
widely acceptable for diagnostic purposes. At present the only diagnostic
methods which are accepted in those countries where aquatic animal health
control programmes are implemented, are based either on isolation of the
pathogen followed by its specific identification, or on demonstration of
pathogen-specific antigens using an immunological detection method.
Mollusc diseases differ in some ways from fish diseases with respect to the
considerations mentioned above. General information on diagnostic techniques
for mollusc diseases is given in Chapter 15.
As explained in Part 3 of the Code, the list of notifiable diseases of aquatic
animals includes only major diseases of proven aetiology and limited
geographic range. The OIE Fish Diseases Commission therefore recommended
the creation of a list entitled 'Other significant diseases'. The diseases on this
list include:
•
those which are serious, but have a broad geographic distribution;
•
those causing significant mortality, transmissible, and of limited
geographic range, but for which the aetiological agent has not yet been
identified;
viii
•
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
those with the potential for causing large losses, but which are too new for
the geographic range to be defined or for the essential epizootiological
elements to be understood.
It is expected that the diseases on this list will either be elevated to notifiable
status or dropped from the list as new information is obtained.
The Manual includes descriptions of diagnostic methods for these 'Other
significant diseases' as well as for the notifiable diseases.
ix
Abbreviations
ABBREVIATIONS
Ab
antibody
Ag
antigen
AO
acridine orange
BF-2
bluegill fibroblast (cell line)
BKD
bacterial kidney disease
BMN(V)
baculoviral midgut gland necrosis virus
BP
Baculovirus
BSA
bovine serum albumin
BSS
balanced salt solution
ceo
channel catfish ovary (cell line)
CC(VD)
channel catfish (virus disease)
CFA
complete Freund's adjuvant
CHSE-214
chinook salmon embryo (cell line)
penaei
CPE
cytopathic effect
DNA
deoxyribonucleic acid
dNFP
deoxynucleotide
EHN(V)
epizootic haematopoietic necrosis (virus)
ELISA
enzyme-linked immunosorbent assay
EPC
Epithelioma papulosum
ESC
enteric septicaemia of channel catfish
EUS
epizootic ulcerative syndrome
FAT
fluorescent antibody test
FCS
fetal calf serum
FITC
fluorescein
HBSS
Hank's basal salt solution
HEPES
N-2-hydroxyethylpiperazine-N'-2 ethane sulfonic acid
IF
immunofluorescence
IFA
incomplete Freund's adjuvant
IF AT
indirect fluorescent antibody test
Ig
IHHN
infectious hypodermal and haematopoietic necrosis
IHN(V)
infectious haematopoietic necrosis (virus)
IPN(V)
infectious pancreatic necrosis (virus)
ISA
infectious salmon anaemia
cyprini (cell line)
isothiocyanate
immunoglobulin
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
X
KDM-2
kidney disease medium
MAb
monoclonal antibody
MBV
Penaeus monodon-tyye
MEM
minimum essential medium
MOI
multiplicity of infection
NAb
neutralising antibody
NeVTA
nerka virus Towada Lake, Akita and Amori prefecture
OCT
embedding medium for frozen tissue specimens
OMV(D)
Oncorhynchus
OPD
o-phenylenediamine
OWD
oyster velar virus disease
PBS
phosphate buffered saline
PBST
phosphate buffered saline containing Tween
PCR
polymerase chain reaction
PFU
plaque forming units
PIB
polyhedral inclusion body
POB
polyhedral occlusion body
RNA
ribonucleic acid
RSD
red spot disease
SDS
sodium dodecyl sulphate
SJNNV
striped jack nervous necrosis virus
SKDM
selective kidney disease medium
SVC(V)
spring viraemia of carp (virus)
TRIS
tris(hydroxymethyl) aminomethane
VHS(V)
viral haemorrhagic septicemia (virus)
VN
virus neutralisation
YHV
yellowhead virus
YTV
yamame tumour virus
baculovirus
masou virus (disease)
Definitions
xi
DEFINITIONS
The International Aquatic Animal Health Code (companion volume to this
Manual) contains a list of definitions which may be consulted for the meaning
of terms used in this Manual. Some terms which are not used in the Code but
which appear in the Manual, are defined below:
Fry
means newly hatched fish larvae.
Sensitivity
means the proportion of true positive tests given in a diagnostic
test, i.e. the number of true positive results divided by the number
of true positive and false negative results.
Specificity
means the probability that absence of infection will be correctly
identified by a diagnostic test, i.e. the number of true negative
results divided by the number of true negative and false positive
results.
Definitions
xiii
ACKNOWLEDGMENTS
The OIE warmly thanks the following persons who contributed to the writing of
the chapters in this Manual:
Dr D. Alderman, Fish Disease Laboratory, MAFF, Barrack Road, The Nothe,
Weymouth, Dorset DT4 8UB, United Kingdom
Prof. S.-N. Chen*, Institute of Fishery Biology, National Taiwan University,
No. 1 Roosevelt Road, Section 4, Taipei, Taiwan
Prof. J. Fryer, Oregon State University, Corvallis, Oregon 97331-3804, USA
#
Dr H. Grizel , IFREMER, Laboratoire de pathologie des invertébrés, 17390 La
Tremblade, France
Prof. T. Hâstein*, Central Veterinary Laboratory (former National Veterinary
Institute), P.O. Box 8156 Dep., 0033 Oslo, Norway
Dr B.J. Hill*, Fish Disease Laboratory, MAFF, Barrack Road, The Nothe,
Weymouth, Dorset DT4 8UB, United Kingdom
Dr A. Hyatt, Australian Fish Health Reference Laboratory, c/o Australian
Animal Health Laboratory, P.O. Bag 24, Geelong, Vic. 3220, Australia
Dr P. de Kinkelin, Laboratoire d'Ichtyopathologie, Bât. de Biotechnologies,
Centre de Recherche de Jouy-en-Josas, Domaine de Vilvert, 78350 Jouy-enJosas, France
Dr C. Michel*, Laboratoire de Virologie et d'Immunologie moléculaires,
Centre de Recherches de Jouy en Josas, Domaine de Vilvert, 78352 Jouy en
Josas Cedex, France
Dr B. Munday, University of Tasmania, School of Science and Technology,
P.O. Box 1214, Launceston, Tasmania 7250, Australia
Dr T. Nakai, Faculty of Applied Biological Science, Hiroshima University,
Higashi-Hiroshima 724, Japan
Dr R.J. Roberts, Department of Aquaculture, University of Stirling, Stirling
FK9 4LA, Scotland, United Kingdom
Dr R. Subasinghe, Food and Agriculture Organization of the United Nations,
Fisheries Department, Room NF 514, Viale délia Terme di Caracalla, 00100
Rome, Italy
xiv
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Dr R Whittington, Elizabeth Macarthur Agricultural Institute, PMB 8,
Camden NSW 2570, Australia.
Dr J. Winton*, Fish and Wildlife Service, National Fisheries and Research
Center, Building 204, Naval Station, Seattle, Washington 98115-5007, USA
Miss G.S. Townshend is thanked for her excellent editorial work.
*
#
Member of the OIE Fish Diseases Commission, 1991 to present
Observer at OIE Fish Diseases Commission meetings, 1991 to present
General information
1
DISEASES OF FISH
CHAPTER 1
GENERAL INFORMATION
A
- GENERAL BASES
OF
FISH HEALTH SURVEILLANCE/CONTROL
1.
PROGRAMMES
T A R G E T P A T H O G E N S AND DISEASES
Target pathogens and fish diseases are included in the Code according to the
following basic considerations: they resist or respond poorly to therapy, have
a restricted geographic range and are of high socio-economic importance,
regardless of the host they infect. The list of fish diseases considered for
notification and certification is currently restricted to five viral diseases.
They are:
Epizootic haematopoietic necrosis (EHN)
Infectious haematopoietic necrosis (IHN)
Oncorhynchus masou virus disease (OMVD)
Spring viraemia of carp (SVC)
Viral haemorrhagic septicaemia (VHS)
2.
O V E R A L L A P P R O A C H F O R A N I M A L H E A L T H C O N T R O L IN FISH C U L T U R E
A comprehensive approach for animal health control in fish culture requires:
-
Assessment of health status of animals in a production site based
upon inspections and standardised sampling procedures followed by
laboratory examinations conducted according to instructions given in
this Manual.
-
The constraint of restocking open waters and farming facilities only
with products having a health status higher than or equal to that of
animals already living in the considered areas.
-
Eradication of disease when possible, by slaughtering of infected
stocks, disinfection and restocking with pathogen-free fish.
-
Notification by every Member Country of its particular requirements,
beside those provided by the Code, for importation of aquatic animals
and animal products.
2
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
If the above procedures are followed, it becomes possible to give adequate
assurance of the health status of aquaculture products for specified diseases,
according to their country, zone or production site of origin.
The issue of a health certificate by the appropriate official, based on a health
status report and examinations in aquatic animals, provides assurance that
the aquaculture products in a defined consignment originate from a whole
country, a zone or a farm/harvesting site free of one or more of the specified
diseases listed in the Code and possibly of other specified diseases (see
model of international certificate in the Code).
The assessment of the health status of fish stocks is based upon inspection of
fish production sites and further laboratory examination of fish organ
samples originating from fish specimens taken among the stock of a defined
fish population. This endeavour requires the fish sample collecting to be
done according to defined sampling size charts and the organ processing to
be conducted according to accepted methods.
B - SAMPLING PROCEDURES
1.
C O L L E C T I O N O F FISH S P E C I M E N S
Two situations can be encountered when collecting fish during inspection of
fish production sites:
- fish exhibit the clinical signs of one of the diseases listed in the Code
or other diseases.
- fish appear clinically normal.
The goals of the inspection/sampling procedures can thus be different: they
are conducted:
-
either to demonstrate the health status of a fish production site,
-
or to confirm that a certain status is being maintained once it has
been achieved after a minimum period of two years of
implementation of the fish surveillance programme enforced in the
country.
1.1. Clinically infected fish
A minimum number of ten moribund fish or ten fish exhibiting clinical
signs of the diseases in question, must be selected and taken. Fish
should be alive when collected. They should be sent to the laboratory
alive or sacrificed and packed separately in sealed aseptic refrigerated
containers or on ice. The freezing of collected fish must be strictly
General information
3
avoided. However, it is highly preferable and recommended to collect
organ samples from the fish immediately after they have been selected
at the fish production site and to store and process the samples as
described in Sections 2 and 3. A label of identification mentioning the
place and time of sampling must be attached to the sample.
1.2. Asymptomatic fish (healthy fish)
Fish collection must encompass a statistically significant number of
specimens, but it is obvious that failure to detect certain pathogens
from the sample does not guarantee the absence of these agents in the
specimen examined or in the stock. This is particularly true of freeranging or feral stocks from which it is difficult to collect a
representative and random sample. However, the risk of a pathogen
escaping the surveillance system is reduced in fish farms whose fish
stocks have been inspected and checked for pathogens for several years
(at least two), insofar as they are not exposed to possible
recontamination by migratory fish.
When a given fish production site harbours a broodstock, it is essential
for one of the sample collections made each year to be focused on the
sexual products (sperm and ovarian fluid) released by broodfish at time
of spawning (see below). If an adult broodstock includes fish of
different ages, the older fish should be selected for sampling:
•
Samples must comprise all susceptible species on the site, with each
lot of a species being represented in the sample group. A lot is
defined as a group of the same fish species that shares a common
water supply and which originate from the same broodfish or
spawning population. For closed-water fish farming such as pond
culture of cyprinid fish, the fish population from a pond with no
water connection with others, constitutes a lot. For closed-water fish
production units which comprise the storage of fish in holding
facilities after pond harvesting and fish grading and sorting, the
fish stock harboured in such a facility can be considered as a lot
providing fish sub-samples are taken from populations of all the
holding tanks.
The geographic origin of samples should be defined by the name of
the sampling site associated with either its geographic co-ordinates
or its localisation along a river course expressed as a kilometric
point.
•
If any moribund fish are present in the fish population to be
sampled, they should be selected first for sample collection and the
remainder of the sample is to be made with randomly selected live
fish from all containers which represent the lot being examined.
4
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
•
The minimum sample size for each lot must be in accordance with
a plan which provides 9 5 % confidence that infected specimens will
be included in the fish sampled, assuming a minimum prevalence
of infection equal to or greater than 2%, 5% or 10%. The minimum
sample size for lots varying from 50 to infinity in size, for each
inspection is given in Table I (23).
•
As in the case of clinically infected fish, organ and fluid samples
must be taken and processed as soon as possible after fish specimen
collection. Sample freezing must be avoided.
Table I
Sample size based on assumed pathogen prevalence in lot
At 2 % prevalence
size of sample
Lot size
50
100
250
500
1,000
1,500
2,000
4,000
10,000
100,000or more
50
75
110
130
140
140
145
145
145
150
(After Ossiander and Wedemeyer.
At 5% prevalence
size of sample
At 1 0 %
prevalence size of
sample
35
45
50
55
55
55
60
60
60
60
20
23
25
26
27
27
27
27
27
30
¡973)
1.3. Sampling specifications according to the objectives of a given fish
surveillance programme
a) Achievement
of the health status of a fish stock/population
given inspection site
•
at a
A fish culture unit must be inspected twice a year during two
years at the appropriate lifestage of the fish and at times of year
when temperature and season offer the best opportunity for
observing clinical signs and isolating the pathogens if present.
Each time, fish must be collected in order to detect a prevalence
of infection equal to or higher than 2 % at 9 5 % confidence level.
Most often, 150 fish will thus be collected on each occasion or,
during one of the two inspections, 150 ovarian fluid samples
will be taken from broodfish if present in the given fish culture
unit.
General information
5
•
If fish health surveillance is focused on wild fish populations at
a given site of inspection or on rearing ponds without holding
facilities in which different fish crops may be pooled, collection
of 150 fish specimens must be done once a year for two years.
Insofar as it is possible specimens of the oldest fish and/or
ovarian fluid must be collected as a priority.
•
During this two-year period, the fish production unit may only
be restocked with fish from a unit whose health status has
already been approved.
b) Maintenance
of the health
status
•
Once a fish production unit including pond fish production units
equipped with holding facilities, has been recognised to be free of
all or certain diseases listed in the Code after two years of
surveillance with laboratory tests and in the absence of any suspect
clinical signs, twice-yearly inspections must continue. However,
collection of fish specimens may be reduced to 30 fish, including
broodfish when available. Moribund fish observed during
inspection visits must, however, be collected for further laboratory
examination.
•
Maintenance of health status of wild fish populations relevant to
diseases listed in the Code at a given site of inspection, can only be
ascertained by annual collection of 150 individuals including as
many broodfish as possible.
•
The fish production unit may only be restocked with fish having a
health status higher than or equal to that of those already present.
•
If, during monitoring of samples, a cytopathic effect (CPE) appears
in cell cultures inoculated with dilutions of the samples being
tested, virus identification procedures have to be undertaken
immediately (see the relevant chapters). Provisions have to be taken
to suspend the approved health status of the production unit and/or
the zone (if it was approved previously) from which the virus
positive sample originated. The suspension of approved status will
be maintained until it is demonstrated that the virus in question is
not the one referred to in the granting of free status.
The above sampling specifications for the achievement and
maintenance of the health status of fish at given fish production sites
imply that all provisions given in Section A.2 (Overall approach for
animal health control in fish culture) are in force.
6
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
2.
S A M P L E M A T E R I A L T O B E USED IN V I R A L AND B A C T E R I O L O G I C A L T E S T S
Sample material depends both on the size of animals and the objective of
testing i.e. diagnosis of overt disease or detection of fish that are
asymptomatic pathogen carriers.
2.1. Specifications according to fish size
Alevin and sac fry: sample entire fish but remove the yolk sac if
present.
Fish 4-6 cm: take the entire viscera including kidney. A piece of
encephalon can be obtained after severing head at level of the rear edge
of operculum and pressing it laterally.
Fish over 6 cm: take the kidney, spleen and encephalon.
Broodfish: take ovarian fluid and/or tissues as described in Chapters 26.
2.2. Specifications according to clinical status
In the case of clinical infection, beside whole alevin or entire viscera,
organs to be sampled are anterior kidney, spleen and encephalon for
virus screening and kidney and spleen for bacterial screening. Samples
from 10 diseased fish will thus be taken and combined to form pools of
a maximum of 5 fish each. The amount of material should not exceed
1.5 g/pool of material from 5 fish.
For detecting asymptomatic carriers, samples may be combined to form
a pool (no more than 5 fish/pool), for a total weight of about 1.5 g.
Pools of ovarian fluid from 5 broodfish must not exceed a total volume
of 5 ml, i.e. 1 ml/broodfish. These ovarian fluid samples are to be taken
individually from every sampled female and not after the pooling of
ova.
Once aseptically removed from fish, organs and/or ovarian fluid
sampled are each split into two parts, one intended for virological
examination, the other for bacteriological examination. For
bacteriological sampling, live fish, newly dead (chilled) fish or
inoculated required medium are preferred.
3.
G E N E R A L PROCESSING
EXAMINATION
OF
ORGANS/FLUID
SAMPLES
FOR
VIROLOGICAL
3.1. Transportation and antibiotic treatment of samples
Pools of organs or of ovarian fluids are placed in sterile vials and
stored at 4°C until virus extraction is performed at the laboratory.
Virus extraction should optimally be done within 24 h after fish
General information
7
sampling but is still acceptable for up to 48 h.
Organ samples may also be transported to the laboratory by placing
them in vials containing cell culture medium or Hanks' basal salt
solution (HBSS) with added antibiotics to suppress the growth of
bacterial contaminants (1 vol. of organ in at least 5 vol. of
transportation fluid). Suitable antibiotic concentrations are: gentamycin
1,000 ug/ml or penicillin (800 IU/ml) and dihydrostreptomycin
(800 ug/ml). The antifungal compounds Mycostatin® or Fungizone®
may also be incorporated into the transport medium at a final
concentration of 400 IU/ml. Serum or albumen 5-10% may be added to
stabilise the virus if the transport time will exceed 12 hours.
3.2. Virus extraction
•
Conduct this procedure below 15°C and preferably between 0°C
and 4°C.
•
Decant antibiotic-supplemented medium from organ sample.
•
Homogenise organ pools with mortar and pestle or electric blender
until a paste is obtained. This must be resuspended in transport
medium to a final dilution of 1:10.
•
If organ samples have not been treated with antibiotics prior to
homogenisation, organ homogenates are to be resuspended in
antibiotic-supplemented medium and incubated in this medium for
2-4 hours at 15°C or overnight at 4°C.
•
Clarify diluted homogenates by centrifugation at 2,000 xg for 15
min. and collect supernatants.
•
Ovarian fluid samples are to be centrifuged in the same way as
organ homogenates and their supernatants used directly in
subsequent steps.
3.3. Treatment to neutralise IPN viruses
In some countries, fish are often asymptomatic carriers of aquatic
birnaviruses (such as infectious pancreatic necrosis virus, IPNV),
which induce a cytopathic effect (CPE) in susceptible cell cultures and
thus complicate and retard isolation and further identification of other
viruses. In such situations, the infectivity of birnaviruses possibly
present in sample homogenate supernatants must be neutralised before
testing for the viruses listed in the Code. However, when it is important
to determine whether IPNV is present, samples must be tested with and
without neutralising antibodies present.
To neutralise birnaviruses, mix equal volumes (200 ul) of a solution of
neutralising antibodies (NAb) to the indigenous birnavirus serotypes of
8
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
IPNV and of the supernatant to be tested. Allow the mixture to react
for one hour at 15°C prior to inoculation onto susceptible cell
monolayers. The titre of the NAb solution used (it may be a multivalent
serum) should be at least 2,000 in a 5 0 % plaque reduction test versus
the IPNV serotypes present in the given geographic area.
When samples are from a country, region, fish population or
production unit considered free from birnavirus infection, this
treatment of the organ homogenate should be omitted.
4.
G E N E R A L PROCESSING O F SAMPLES INTENDED FOR
EXAMINATION
BACTERIOLOGICAL
As in viral infections, internal organs may be used as a source of isolation
whenever systemic infection is suspected. However, active proliferation of
saprophytic associated microorganisms is such a disadvantage that live fish
are preferred for bacteriological examination. The fact that no antibiotic
substances may be added to the transport medium in which the samples are
collected reinforces this preference.
Renibacterium
salmoninarum
is an instance of these cases in which
biological products can be used with some advantage. Sexual products,
namely the ovarian fluid, seem to represent quite a good source of
pathogenic bacteria in mature salmonids.
C - M A T E R I A L S AND B I O L O G I C A L P R O D U C T S R E Q U I R E D
F O R T H E I S O L A T I O N AND I D E N T I F I C A T I O N O F F I S H P A T H O G E N S
1.
F I S H VIRUSES
1.1. Fish cell lines
The following four fish cell lines will be required to test for the fish
pathogens mentioned in the Code:
Bluegill fibroblast (BF-2)
Channel catfish ovary (CCO)
Chinook salmon embryo (CHSE-214)
Epithelioma papillosum cyprini (EPC)
Technical information on the use of these cells for the isolation of the
fish pathogens listed in the Code is given in Table II.
1.2. Culture media
Traditional Eagle's minimum essential medium (MEM) with Earle's
Salt supplemented with 10% fetal calf serum, antibiotics and 2 mM
L-glutamine is the most widely used medium for fish cell culture.
General information
9
Table II
Technical information on the most suitable fish cell lines for detection of
the viral agents listed in the OIE Code
Cell line nomenclature
BF-2
ceo
CHSE-214
EPC
Cell morphology
Fibro­
blastic
Fibro/
epithelioid
Epitheli­
oid
Epithelioid
Temperature range (°C)
15-28
15-35
4-25
10-33
20
30
20
30
20
35
16
30
150
300
150
300
8
5-6
10
8
Properties
Culture characteristics
Optimum growth temp. (°C)
4
2
Inoculum (No. cells xl0 /cm )
(1)
Saturation density (No. cells x
10 /cm )
4
2
Time to saturation (days) (2)
( 1 ) To achieve loose confluency within 24 hours. (2) At optimal growth
temperature.
However, Stoker's medium (Stoker and McPherson, 1961) which is a
modified form of the above medium comprising a double strength
concentration of certain amino acids and vitamins, is particularly
recommended
to
enhance
cell
growth,
using
the
same
supplementations as above plus 10% tryptose phosphate.
These media are buffered with either sodium bicarbonate, 0.16 M
trihydroxymethyl aminomethane (Tris) HC1, or, preferably, 0.02 M N2-hydroxyethylpiperazine-N'-2-ethane sulfonic acid (HEPES). The use
of sodium bicarbonate alone is restricted to those cell cultures made in
tightly closed cell culture vessels.
For cell growth, the fetal bovine serum content of the medium is
usually 10%, whereas for virus isolation or virus production it may be
reduced to 2%. Similarly the pH is 7.3-7.4 for cell growth and adjusted
to 7.6 for virus production or virus assay.
The composition of the most frequently used antibiotic mixture is
penicillin 100 IU/ml and dihydrostreptomycin 100 ug/ml. Mycostatin
(50 IU/ml) may be used if a fungal contamination is expected. Other
antibiotics or antibiotic concentrations may be used as convenient for
the operator depending on the antibiotic sensitivity of the bacterial
strains encountered.
10
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
1.3. Virus positive controls and antigen preparation
a) Virus
nomenclature
•
Epizootic haematopoietic necrosis v i m s (EHNV)
•
Infectious haematopoietic necrosis virus (IHNV)
•
Oncorhynchus masou virus (OMV)
(Syn. Salmonid herpesvirus type 2)
•
Spring viraemia of carp virus (SVCV)
•
Viral haemorrhagic septicaemia virus (VHSV)
(Syn. Egtved virus)
b) Virus
production
For the production of most of these viruses the susceptible cell
cultures must be inoculated with fairly low multiplicities of
infection (MOI) i.e. 10" to 10" PFU per cell. However, the MOI
for OMV has to be increased up to 0.1-1 PFU/cell. Furthermore, the
best yields for OMV are obtained using inocula containing cell
debris of the OMV-infected previous cell monolayer.
2
c) Virus preservation
•
3
and storage
Dilute virus-containing cell culture fluids in order to obtain
virus titres averaging 1-2 x 10 PFU/ml.
6
•
Dispense the resulting viral suspensions into sterile vials at
volumes of 0.3-0.5 ml each.
•
Freeze and store each series of reference virus stocks at -80°C,
and check the titre of each virus stock every two months if it has
not been used during that time interval.
Lyophilisation:
long term storage (decades) of the seeds of
reference virus strains is achievable by lyophilisation. For this
purpose, viral suspensions in cell culture medium supplemented
with 10% fetal calf serum (FCS) are mixed vol./vol. with a 5 0 %
sodium glutamate solution in distilled water before processing.
Seal or plug under vacuum and store at 4°C, in the dark.
2.
F I S H BACTERIA
2.1. Culture media
Few species of fish pathogenic bacteria require special media for
cultivation, and most of usual isolation peptones (trypsin-soya, brainheart, etc.) can conveniently be used. However, the low optimal
temperatures of some species result in slow growth and mean that
General information
11
small colonies are frequently obtained during isolation. In these cases it
is usual to add enrichment factors such as serum or blood at 5-10% in
order to improve the cultivation. Similarly, rapid diagnostic systems
are convenient in theory, but in practice it is wise to question the
relative value of negative characteristics, which may result merely from
insufficient metabolic activity.
Conversely Renibacterium salmoninarwn is a very fastidious organism
and requires special media enriched with cysteine (No. A l IB).
Bacteriology of fish is generally conducted at temperatures between 20
and 26°C. It is sometimes necessary or useful to have access to several
incubators. R. salmoninarwn,
Flexibacter psychrophilus
and others
need 15°C for optimal growth and many of the bacteria isolated from
warm water fish may be studied at 30 or 37°C to accelerate the
diagnostic steps.
2.2. Conservation
Bacterial strains can be conserved in the short term on ordinary media,
placing the slants or broths at 4°C. For most strains, the use of
commercial conservation media or agar slants with mineral oil will
extend viability to 1-2 years under the same conditions without further
special requirements.
Freezing is probably the best way to preserve bacterial suspensions of
high titre. However, it does not always prevent some phenotypical
characteristics from changing. When stability of characteristics such as
virulence is a major purpose it may be better to use lyophilisation,
although the number of viable bacteria may be decreased dramatically.
Different kinds of supports have been proposed to improve the efficacy
of freezing and lyophilisation, namely glycerol 5 to 15% in the first
case, skim milk, lactose, dextran 5 to 10% in the second one. There is
no general rule, and convenient conditions have to be determined in
prior trials for all species. The addition to fresh cultures of one volume
of support containing Bactopeptone 1 1 % + Dextran 4 % has provided
excellent results for lyophilisation of certain fish bacteria, but other
formulas would be worth testing in many cases.
3.
SEROLOGY
3.1. Production of rabbit antisera and polyclonal antibodies to fish
viruses
There are various ways in which antibodies against fish viruses can be
raised in rabbits. However; titre and specificity are influenced by the
inoculation programme used (see Hill et al., 1981). The following
immunisation protocols may be used to produce antisera for use in the
virus isolation and/or identification procedures described later.
12
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
a) Antisera to IPNV
Intravenous injection with 50-100 ug of purified virus on day 0
followed by an identical booster on day 2 1 . Bleeding 5-7 days later.
Rabbits may be reused if not bled completely.
b) Antisera to other
viruses
The immunisation protocols alternate an intramuscular
intradermal injection with further intravenous boosters:
or
Day 0, primary injection: 500-1,000 ug of virus are mixed with
adjuvant (Freund's complete or other ) vol./vol. giving a total
volume of 1.2 ml. This antigen is delivered to the rabbit as
multipoint intradermal injections (20 points on each side) after the
animal has been shaved.
1
Day 21: collect about 20 ml of blood and check for reactivity
(neutralisation, fluorescence) and boost intravenously with the same
amount of virus as in the primary injection.
Prior to the intravenous booster injection, the rabbit has to be
treated with promethazine (12 mg IM) to prevent possible
anaphylactic response.
Day 28: sample the blood, check the serum reactivity and bleed or
boost according to the results.
3.2. Antisera to fish bacteria
It is still difficult to obtain antimicrobial sera in large amounts from
commercial sources, and it will be often necessary to prepare such
antisera. The general methods are the same as for viruses. Bacterial
antigens are frequently used as crude preparations killed by heating or
formalin 3.5 p. 1000. Rabbit injection at increasing doses can be
intramuscular, every week, with adjuvant the first time, or intravenous
at 3-4 day intervals without adjuvant. A booster injection is often
required after 15 days.
A special multipoint intradermal schedule has proven very efficient for
anti-Renibacterium
sera production, and is also valuable for other
poorly antigenic bacteria.
The antigen is heat killed (60°C, 45 min) and adjusted to 2 mg/ml. The
flanks of the animals are thoroughly shaved, and multipoint
intradermal injections are performed using total amounts of 1 mg
bacteria/animal, according to the following schedule:
Use of Freund's adjuvants may be restricted on animal welfare grounds.
Alternative synthetic adjuvants include trehalose dimycolate (TDM) and
monophosphate lipid A (MPL).
General information
13
bacteria/animal, according to the following schedule:
Day
Day
Day
Day
Day
1
21
42
63
68
1 mg + complete Freund adjuvant (CFA) v/v
1 mg + incomplete Freund adjuvant (IFA) v/v
1 mg + IF A
1 mg antigen
Collecting of blood samples (about 30 ml)
Withdrawal injections and bleeding for serum collection may be
repeated at one month intervals.
3.3. Processing and storage of immune sera
After blood clotting, collect and centrifuge the serum at 20°C and heat
it for 30 min at 56°C. Filter the resulting heat-inactivated serum
through a 450 nm membrane and temporarily store it at +4°C during
the time necessary for the screening of its reactivity and specificity and
for checking that these properties are not affected by preservation
conditions (e.g. freezing or lyophilisation). Sterile rabbit sera can be
kept for at least two months at 4°C without any change in their
properties.
Dispense (usually as small volumes) and freeze at -20°C or lyophilise.
Immunoglobulins may be extracted from antisera using conventional
methods suitable for Ig purification. Selective attachment to protein A
constitutes a reliable and effective method. The concentration of Ig
solutions is adjusted to the values required for further conjugate
preparation or storage.
Preservation of Ig: Mix vol./vol. a solution of Ig of titre 2 mg/1 with
sterile pure glycerol and keep at -20°C. Ig of solutions of higher titre
may also be prepared in glycerol.
3.4. Mouse monoclonal antibodies to fish viruses and bacteria
Monoclonal antibodies to several fish viruses (IPNV, VHSV, IHNV,
CCV) have been raised during the past years. Some of them, singly or
as two or three associated MAbs, have given rise to biological reagents
suitable for the identification of virus groups (IPN, VHS, IHN). Other
MAbs, taken individually or as components of Ab panels, allow
accurate typing of VHSV and IHNV. These MAbs can be obtained
from the Reference Laboratories listed at the end of the Manual.
The production of monoclonal antibodies to bacteria has also been
described. It has resulted in the development of commercial diagnostic
kits for Renibacterium in 1992, but in most cases remains limited to
specialised laboratories.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
14
may be impaired by processes such as enzymatic or radio-labelling or
lyophilisation. It is thus mandatory to test the stability of every MAb to
processing and preservation conditions.
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herpesvirus of Ictalurid fish. J. Virol., 8(4), 525-533.
62.
W O L F K. & QUIMBY M . C . (1973). - Fish viruses: Buffers and methods for
plaquing eight agents under normal atmosphere. Appl. Microbiol., 25,
659-664.
63.
W O L F K. (1988). - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 476 p.
64.
WUNNER W . H . & PETERS D . (1991). - Family Rhabdoviridae. In: Francki
R.I., Fauque C M . , Knudson D . L . , Brown F. Classification and
nomenclature of viruses. Archives of Virology. Supplementum 2. Springer,
New York, Vienna, 250-262.
Epizootic haematopoeitic necrosis virus
21
DISEASES OF FISH
DISEASES NOTIFIABLE TO THE OIE
CHAPTER
2
EPIZOOTIC HAEMATOPOIETIC NECROSIS
(B413)
SUMMARY
Epizootic
of redfin
mykissj.
restricted
condition,
haematopoietic necrosis (EHN) is an iridovirus
infection
perch (Perca fluviatilis^ and rainbow trout fOncorhynchus
The geographic
range of the infection is
currently
to A ustralia. For a recent and more detailed review of the
see Wolf (10).
The infection offish is most often lethal in perch and much less so in
rainbow trout, in a clinical context of haemorrhages and oedema
accompanied by necrotic lesions of the vascular walls, liver, spleen
and haematopoietic tissue of the kidney. Necrotising hepatitis seems
to be a consistent sign of the condition.
The antigenicity of EHNV is currently under investigation
using
rabbit antibodies (Ab). This virus has not so far generated
synthesis
ofNAb but can be identified by immunofluorescence
or ELISA tests.
However, indirect fluorescent
antibody tests (IFA) and Westernblotting indicate that EHNV shares common antigenic domains with
two other fish iridoviruses isolated from the sheatfish ('Silurus
glanisj and the catfish (Zctalurus melase in Europe and also with frog
virus 3 and Bohle iridovirus.
The epidemiology
of EHNV in rainbow trout is
incompletely
understood. Infection may recur annually at a production site but it
is uncertain whether this is due to persistence of infection on the site
or reintroduction of infection from wild fish in the catchment. A
carrier state in naturally infected rainbow trout appears to be very
uncommon. Neither EHNV antigen nor anti-EHNV antibody are
detected in rainbow trout surviving an outbreak of EHNV.
The factors modulating the susceptibility of fish to EHNV infection
are poorly understood. Clinical disease is associated with poor
water quality. In rainbow trout infection occcurs naturally from 1117°C and experimentally from 8-21°C. Infection in redfin perch is
non-permissive
at temperatures below 12°C. The following
fish
species were found to be susceptible to EHNV following
bath
exposure: redfin perch, rainbow trout, Macquarie perch (Macquaria
australasica), mosquito fish (Gambusa affinis), silver
perch
22
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
(Bidyanus bidyanus) and mountain galaxias (Galaxias olidus).
Both juvenile and adult redfin perch may be affected in
but juveniles may be more susceptible to the disease.
outbreaks,
The diagnostic procedures for EHNV are based upon isolation of
virus in cell culture, ELISA, IFA, and electron microscopy.
Antigen
capture ELISA has a sensitivity of 80% and a specificity of 99%
when used to test tissues from both rainbow trout and redfin perch.
Antigen capture ELISA is the method of choice for confirming the
cause of CPE in cell culture; however, IFA and electron
microscopy
are also useful. IFA or immunoperoxidase staining may be used also
for diagnosis on formalin fixed tissues..
DIAGNOSTIC PROCEDURES
The diagnosis of EHN is based upon direct methods which are either the
isolation of EHN virus (EHNV) in cell culture followed by its immunological
identification (conventional approach), or the immunological demonstration of
EHNV antigen (Ag) in infected fish tissues.
Due to insufficient knowledge of the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been recognised as a
valuable diagnostic method for assessing the virus status of fish populations.
However, the validation of some serological techniques for diagnosis of certain
fish virus infections could arise in the near future, rendering the use of fish
serology more widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
during overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
kidney, spleen and liver.
-
during dormant infection (detection of asymptomatic virus carrier fish):
encephalon (any size fish) and/or ovarian fluid from broodfish at time of
spawning.
Sampling procedures: see Chapter 1 Part B.
1.
STANDARD MONITORING METHODS FOR EHNV
1.1. Isolation of EHNV in cell culture
Cell line(s) to be used: BF-2.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10
organ
23
Epizootic haematopoeitic necrosis virus
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-old BF-2 cell
monolayers. Inoculate at least 2 c m of cell monolayer with
100 pi of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 10-15°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 20-22°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, during 14 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation phase. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the tested homogenate
supernatants, identification procedures have to be undertaken
immediately (see below). If a fish health
surveillance/control
programme is being implemented, provisions have to be taken
to suspend the approved health status of the production
unit
and/or the zone (if it was approved previously) from which the
virus positive sample originated. The suspension of approved
status will be maintained until it is demonstrated that the virus
in question is not EHNV.
iv)
If no CPE occurs, except in positive control cell cultures,
subcultivation steps have to be made even after 7 days of
incubation, in certain of the infected cell cultures. However, if
no CPE is observed even in positive controls, another series of
virological examinations have to be undertaken, using
susceptible cells and new batches of organ samples.
c) Subcultivation
i)
procedures
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
24
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
ii)
Centrifuge at 2,000 x g for
supernatant.
15 min at 4°C and
collect
iii)
Repeat optional neutralisation test to IPNV if needed, with
dilution of the above supernatant (1:1 to 1:100).
iv)
Inoculate BF-2 cell monolayers as described above (1.1.a).
v)
Incubate and monitor as in 1.1 .b.
1.2. Virus identification
a) Neutralisation
test
EHNV cannot be identified by neutralisation as the antisera
generated by the immunisation of rabbits have few neutralising
antibodies.
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
2
i)
Prepare monolayers of BF-2 cells in 2 c m wells of cell culture
plastic plates or on coverglasses in order to reach around 8 0 %
confluency, which is usually achieved within 4 hours of
incubation at 25°C (seed 6 cell monolayers per virus isolate to
be identified, plus 2 for positive and 2 for negative controls).
The FCS content of the cell culture medium can be reduced to
2-4%. If numerous virus isolates have to be identified, the use
of Terasaki plates is strongly recommended.
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of EHNV in a similar way,
in order to obtain a virus titre of about 5,000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 20°C for 24 hours.
v)
When the incubation time is over, aspirate the cell culture
medium, rinse once with PBS 0.01 M pH 7.2, then 3 times
briefly with cold fixative. This fixative will be acetone (stored
at -20°C) for coverglasses or a mixture of acetone 30%ethanol 7 0 % (vol./vol.), also stored at -20°C.
25
Epizootic haematopoeitic necrosis virus
vi)
Afterwards, let the fixative act for 15 min. A volume of 0.5 ml
is adequate for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to EHNV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the dried cell monolayers by 4 rinsing steps with
the above PBS and eliminate this buffer thoroughly after the
last rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber. The volume of solution to be used
is 0.25 ml/2 c m well.
2
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Observe the treated cell monolayers on plastic plates
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
xv)
Conduct this observation under incident UV light using a
microscope with xlO eye pieces and x20 to x40 objective lens
having numerical aperture >0.65 and >1.3 respectively.
Positive and negative controls must be found to give the
expected results prior to any other observation.
c) Enzyme-linked
immunosorbent
assay (ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for EHNV, in PBS 0.01 M pH 7.2 (200
ul/well). Ig may be polyclonal or monoclonal Ig originating
most often from rabbit or mouse, respectively.
ii)
Incubate overnight at 4°C.
26
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 ul/well).
v)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vi)
Rinse 4 times with PBST.
vii) Dispense 100 ul/well of a 2 or 4 step dilution of the virus to be
identified and of EHNV control virus, and allow to react with
the coated antibody to EHNV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
ix)
Add to the wells, biotinylated polyclonal antibody to EHNV.
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(O-phenylenediamide, OPD or other approved chromogen).
Stop the course of the test when positive controls react, and
read the results.
xv) Alternatively: add H2O2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2.
DIAGNOSTIC PROCEDURES FOR CONFIRMATION
OUTBREAKS
O F EHN
IN S U S P E C T E D
Confirmation of EHNV can be achieved by any of the following methods.
2.1. Conventional
virus
isolation
identification as in Section 1.
with
subsequent
2.2. Virus isolation with simultaneous identification
Not possible; see 1.2.a.
serological
Epizootic haematopoeitic necrosis virus
27
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom of
the wells of a plastic cell culture plate.
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required for
virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in 1.2.b.
points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and block
with 5 % skim milk or 1% bovine serum albumin (BSA), in PBST
for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to EHNV and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Reveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in 1.1.
2.4.
ELISA
a) Microplate
processing
As 1.2.c of this chapter up to point iv (inclusive).
b) Sampling
procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection of fish specimens
B.2.1. & 2.2. for the selection of materials sampled.
c) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ
homogenates.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
28
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol./vol.) as 1.2.c point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.c.
REFERENCES
1.
AUBERTIN A . M . ( 1 9 9 1 ) . - Family Iridoviridae. In: Francki R.J., Fauque
C M . , Knudson D . L . & Brown F. Classification and nomenclature of
viruses. Archives of Virology, Supplementum 2 . Springer, New York,
Vienna, 1 3 2 - 1 3 6 .
2.
HEDRICK R . P . , M C D O W E L L T . S . , A H N E W . , T O R H Y C. & D E KINKELIN P.
( 1 9 9 2 ) . - Properties of three iridovirus-like agents associated with systemic
infections offish. Dis. Aquat. Org., 13, 2 0 3 - 2 0 9 .
3.
HENGSTBERGER
S.G.,
HYATT
A.D.,
SPEARE
R.S.
&
COUPAR
B.E.H.
( 1 9 9 3 ) . - Comparison of epizootic haemopoeitic necrosis virus and Bohle
iridoviruses, recently isolated Australian iridoviruses. Dis. Aquat. Org.,
15, 9 3 - 1 0 7 .
4.
H Y A T T A . D . , E A T O N B . T . , HENGSTBERGER S. & RUSSEL G.
(1991).
-
Epizootic hematopoietic necrosis virus: detection by ELISA, immunohistochemistry and electron microscopy. J. Fish Dis., 14, 6 0 5 - 6 1 8 ) .
5.
L A N G D O N J.S., HUMPHREY J . D . , WILLIAMS L . M . , H Y A T T A . D . , WESTBURY
H.A. ( 1 9 8 6 ) . - First virus isolation from Australian fish: an iridovirus-like
pathogen from redfin perch. Perca fluviatilis L. J. Fish Dis., 9, 2 6 3 - 2 6 8 .
6.
7.
LANGDON J.S. ( 1 9 8 9 ) . - Experimental transmission and pathogenicity of
epizootic haematopoietic necrosis virus (EHNV) in redfin perch, Perca
fluviatilis L., and 1 1 other teleosts. J. Fish Dis., 12, 2 9 5 - 3 1 0 .
WHITTINGTON R . J . & STEINER K.A. ( 1 9 9 3 ) . - Epizootic haematopoietic
necrosis virus (EHNV): improved ELISA for detection in fish tissues and
cell cultures and an efficient method for release of antigen from tissues. J.
Virol. Meth., 43, 2 0 5 - 2 2 0 .
8.
W H I T T I N G T O N R . J . , P H I L B Y A.,
REDDACLIFF G.L.
&
MACGOWN
A.R.
( 1 9 9 4 ) . - Epidemiology of epizootic haematopoietic necrosis virus (EHNV)
infection in farmed rainbow trout, Oncorhynchus
mykiss (walbaum):
Epizootic haematopoeitic necrosis virus
29
findings based on virus isolation, antigen capture ELISA and serology. J.
Fish Dis., 1 7 , 2 0 5 - 2 1 8 .
9.
W O L F K . , G R A V E L L M . & M A L S B E R G E R K G . ( 1 9 6 6 ) . - Lymphocystis virus:
isolation and propagation in centrarchid fish cell line. Science, 151, 1 0 0 4 1005.
10.
W O L F K . ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 pp.
Infectious haematopoietic necrosis
31
CHAPTER
3
INFECTIOUS HAEMATOPOIETIC NECROSIS
(B405)
SUMMARY
Infectious haematopoietic necrosis (IHN) is a rhabdovirus
infection
of rainbow trout (Onchorhyncus mykiss) including
steelhead,
several Pacific salmon i.e. sockeye (O. nerka), chinook (O.
tshawystcha), chum (O. keta), yamame (O. masou), amago (O.
rhodurus), and more recently coho (O. kisutch) and Atlantic salmon
(Salmo salar). Until 1987, the geographic range of IHN was limited
to the North Pacific Rim (North America and the Far East) but it has
recently spread to continental Europe. For a recent and more
detailed review of the condition, see Wolf (7).
IHN has become a real matter of concern because of its clinical and
economic consequences
in trout and salmon farming
and in
fisheries. Infection is often lethal, due to the impairment of the salt­
water balance, which occurs in a clinical context of oedema and
haemorrhages.
Virus multiplication
in endothelial cells of blood
capillaries, haematopoietic tissues and nephron cells, underlies the
clinical signs.
In survivors, IHN virus (IHNV) infection results in a strong
protective immunity, synthesis of circulating antibodies to IHNV as
well as, in certain individuals, an asymptomatic carrier state. This
carrier state frequently
leads to virus shedding via the sexual
products at times of spawning.
On the basis of antigenic studies conducted with
neutralising
polyclonal antibodies from rabbit (rabbit antisera), IHNV isolates
appear to form a homogeneous
virus group. However,
mouse
monoclonal antibodies have revealed at least three
neutralising
virus-types (and more likely five) as well as the existence of an
IHNV-group non-neutralising epitope borne by the nucleocapsid (N
protein). Variations in the virulence of IHNV strains have been
recorded during both natural cases of disease and in experimental
infections.
The reservoirs
of IHNV
are clinically
infected fish
and
asymptomatic carriers from either cultured, feral or wild fish. Virus
is shed via faeces, urine, sexual fluids and external mucus whereas
kidney, spleen, encephalon and digestive tract are the sites in which
virus is the most abundant during the course of overt infection. The
transmission of IHNV is horizontal and possibly vertical or rather,
"egg-associated". Horizontal transmission may be direct or through
32
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
a vector, water being the major abiotic vector. Animate vectors and
fomites also act in IHNV transmission. Egg-associated
transmission
seems to be an infrequent event but is the only
mechanism
substantiating
the onset of IHN in alevins originating
from
disinfected eggs which had been incubated and hatched in virus-free
water. Once IHNV is established in a farmed stock or in a watershed
due to either spawning of infected migratory fish or from river
restocking for recreational purposes, the disease becomes endemic
because of carrier fish.
Beside the salmonid fish species susceptible to natural
IHNV
infections, pike fry (Esox lucius) can be easily infected via the water
route under experimental conditions. As usual, among individuals of
each fish species, there is a high degree of variability
in
susceptibility to IHNV. The age of fish appears to be extremely
important: the younger the fish, the more susceptible to overt
infection. As with VHS, a good overall fish health condition seems to
decrease the susceptibility to overt IHNV, while handling stress and
other types of stress frequently cause subclinical infection to become
overt.
The most prominent environmental factor affecting IHNV is water
temperature. Clinical disease does not occur above I8°C under
natural conditions, but certain virus strains are only
pathogenic
below 14-15°C.
The diagnostic procedures for IHNV are all based upon direct
methods. The conventional approach is the most widely used and
involves isolation of the virus in cell culture followed
by
immunological identification by neutralisation,
immunofluorescence
or ELISA.
Control methods currently rely on the implementation
of control
policy rules and of hygiene practices in the operating of salmonid
husbandry. The thorough disinfection of fertilised eggs and the
incubation of eggs and further rearing of fry and alevins in premises
completely separated from those harbouring possible virus carriers
and free from possible contact with fomites,
are critical for
preventing the occurrence of IHNV in a defined fish production site.
Vaccination is only at an experimental stage at present.
DIAGNOSTIC PROCEDURES
The monitoring for and diagnosis of IHN is based upon direct methods which
are either the isolation of IHN virus (IHNV) in cell culture followed by its
immunological identification (conventional approach), or the immunological
demonstration of IHNV antigen (Ag) in infected fish tissues.
33
Infectious haematopoietic necrosis
Due to insufficient knowledge on the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been accepted as a routine
diagnostic method for assessing the virus status of fish populations. However,
the validation of some serological techniques for diagnosis of certain fish virus
infections could arise in the near future, rendering the use of fish serology more
widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
during overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
kidney, spleen and encephalon.
-
during dormant infection (asymptomatic virus carrier fish): encephalon
(any size fish) and/or ovarian fluid from broodfish at time of spawning.
Sampling procedures: see Chapter 1 Part B.
1.
STANDARD MONITORING M E T H O D S FOR
IHN
1.1. Isolation of IHNV in cell culture
Cell line(s) to be used: EPC.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10 organ
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-old EPC cell
monolayers. Inoculate at least 2 c m of cell monolayer with
100 pi of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 10-15°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 15°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, for 7 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation period. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
34
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the homogenate, identification
procedures must be undertaken immediately (see below). If a
fish health surveillance/control
programme is being imple­
mented, steps have to be taken to suspend the approved health
status of the production unit and/or the zone (if it was
approved previously) from which the suspected vims positive
sample originated. The suspension of approved status will be
maintained until it is demonstrated that the virus in question
is not IHNV.
iv)
If no CPE occurs after 7 days of incubation (except in positive
control cell cultures), subcultivation of the inoculated cell
cultures must be performed. However, if no CPE is observed in
positive controls, another series of virological examinations
must be undertaken, using susceptible cells and new batches of
organ samples.
c) Subcultivation
procedures
i)
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
ii)
Centrifuge at 2,000 x g for
supernatant.
iii)
If required, repeat neutralisation test to IPNV as previously
described, with dilution of the above supernatant (1:1 to
1:100).
iv)
Inoculate EPC cell monolayers as described above (1.l.a).
v)
Incubate and monitor as in l.l.b.
vi)
If no CPE occurs the test may be declared negative.
15 min at 4°C and
collect
1.2. Virus identification
a) Neutralisation
i)
test
Collect the culture medium of the cell monolayers exhibiting
CPE and centrifuge it at 2,000 xg for 15 min at 4°C, or filter
through a 450 run pore membrane to remove cell debris.
Infectious haematopoietic necrosis
35
ii)
Dilute virus-containing medium 10-2
iii)
Mix aliquots (for example 200 pi) of each dilution with equal
volumes of an antibody solution for IHNV, and similarly treat
aliquots of each virus dilution with cell culture medium.
a n c
j irj-4.
(The titre of neutralising antibody (NAb) solution must be at
least 2,000 for 5 0 % plaque reduction.)
iv)
In parallel, a neutralisation test must be performed against
homologous IHNV (positive neutralisation test).
v)
If required, do a similar neutralisation test with antibodies to
IPN, to ensure that no IPNV contaminant might have escaped
the first anti-IPNV test.
vi)
Incubate all the mixtures at 15°C for 1 hour.
vii) Transfer aliquots of each of the above mixtures onto EPC cell
monolayers (inoculate 2 cell cultures per dilution) and allow
adsorption to occur for 0.5 to 1 hour at 15°C. Twenty-four or
12 well cell culture plates are suitable for this purpose, using a
50 pi inoculum.
viii) When adsorption is completed, add cell culture medium
supplemented with 2 % FCS and buffered at pH 7.4-7.6 into
each well and incubate at 15°C.
ix)
Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in 2 0 % ethanol.
x)
The tested virus is identified as IHNV when CPE is prevented
or noticeably delayed in the cell cultures which received the
virus suspension treated with the IHNV-specific antibody,
whereas CPE is evident in all other cell cultures.
xi)
In the absence of any neutralisation by NAb to IHNV, it is
mandatory to conduct an indirect fluorescent antibody test
with the suspect sample. Indeed some cases of antigenic drift
of surface antigen have been observed, resulting in occasional
failure of the neutralisation test using NAb to IHNV.
36
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
2
i)
Prepare monolayers of EPC cells in 2 c m wells of cell culture
plastic plates or on coverglasses in order to reach around 8 0 %
confluency within 24 hours of incubation at 22°C (seed 6 cell
monolayers per virus isolate to be identified, plus 2 for positive
and 2 for negative controls). The FCS content of the cell
culture medium can be reduced to 2-4%. If numerous virus
isolates have to be identified, the use of Terasaki plates is
strongly recommended.
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of IHNV in a similar way,
in order to obtain a virus titre of about 5,000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 15°C for 24 hours.
v)
Remove the cell culture medium, rinse once with PBS 0.01 M
pH 7.2, then 3 times briefly with cold acetone (stored at 20°C) for coverglasses or a mixture of acetone 30%-ethanol
7 0 % (vol./vol.), also at -20°C, for plastic wells.
vi)
Let the fixative act for 15 min. A volume of 0.5 ml is adequate
for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to IHNV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the dried cell monolayers by 4 rinsing steps with
the above PBS-Tween and eliminate this buffer thoroughly
after the last rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber and do not allow evaporation to
occur. The volume of solution to be used is 0.25 ml/2 c m
well.
2
37
Infectious haematopoietic necrosis
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Examine the treated cell monolayers on plastic plates
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
xv) Examine under incident UV light using a microscope with xlO
eye pieces and x20 to x40 objective lens having numerical
aperture >0.65 and >1.3 respectively. Positive and negative
controls must be found to give the expected results prior to any
other observation.
c) Enzyme-linked
immunosorbent
assay
(ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for IHNV. in PBS 0.01 M pH 7.2 (200 ulAvell).
Ig may be polyclonal or monoclonal Ig originating most often
from rabbit or mouse, respectively. For the identification of
IHNV, monoclonal antibodies specific for certain domains of
the nucleocapsid protein (N) are suitable.
ii)
Incubate overnight at 4°C.
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 ul/well).
v)
Rinse 4 times with PBST.
vi)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vii) Dispense 100 ul/well of 2 or 4 step dilutions of the virus to be
identified and of IHNV control virus, and allow to react with
the coated antibody to IHNV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
38
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
ix)
Add to the wells, biotinylated polyclonal antibody to IHNV; or
MAb to N protein specific for a domain different from the one
of the coating MAb and previously conjugated with biotin.
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times with PBST.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(O-phenylenediamide, OPD or other approved chromogen).
Stop the course of the test when positive controls react, and
read the results.
xv) Alternatively: add H2O2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2.
DIAGNOSTIC PROCEDURES
OUTBREAKS
FOR CONFIRMATION
OF I H N
IN
SUSPECTED
Confirmation of IHNV can be achieved by any of the following methods:
2.1. Conventional
identification
virus
isolation
with
subsequent
serological
As Section 1.
2.2. Virus isolation with simultaneous serological identification
a) Sampling
procedures
As B . l . l . in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ homogenates
B.3.3. for treatment to neutralise birnaviruses (if required).
39
Infectious haematopoietic necrosis
c) Virus identification
by neutralisation
test
i)
Dilute organ homogenates 1:100, 1:1,000 and 1:10,000 in cell
culture medium.
ii)
Mix with equal volume of a solution of antibody to IHNV as in
1.2.a, inoculate the cell monolayers, incubate at 15°C and
monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
U.c.
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom of
the wells of a plastic cell culture plate.
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required
for virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in
1.2.b. points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and block
with 5 % skim milk or 1% bovine serum albumin (BSA), in
PBST for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to IHNV and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Reveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in
1.1.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
40
2.4.
ELISA
a) Microplate
processing
As 1.2.C of this chapter up to point iv (inclusive).
b) Sampling
procedures
See the following sections in Chapter 1 (General Information):
B . 1.1. for the selection of fish specimens
B . 2 . 1 . & 2 . 2 . for the selection of materials sampled.
c) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B . 3 . 1 . for transportation
B . 3 . 2 . for virus
homogenates.
extraction
and obtaining
of
organ
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol./vol.) as 1.2.c point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.c.
REFERENCES
1.
A M E N D D . , YASUTAKE W . & M E A D R . ( 1 9 6 9 ) . - A hematopoietic virus
disease of rainbow trout and sockeye salmon. Trans. Am. Fish. Soc, 98,
796-804.
2.
A R N Z E N J . M . , R I S T O W S.S., HESSON C P . & L I E N T Z J . ( 1 9 9 1 ) . - Rapid
fluorescent antibody test for infectious hematopoietic necrosis virus
( I H N V ) utilizing monoclonal antibodies to nucleoprotein and glycoprotein.
J. Aquat. Anim. Health, 3, 1 0 9 - 1 1 3 .
3.
FIJAN N . , SULIMANOVIC D . , BEARZOTTI M . , M U Z I N I C D . , ZWILLENBERG
L.O.,
CHILMONCZYK S., V A U T H E R O T J . F . & D E KINKELIN P . ( 1 9 8 3 ) .
-
Some properties of the Epithelioma papulosum cyprini ( E P C ) cell line
from carp (Cyprinus carpió). Ann. Virol, Institut Pasteur, 134E, 2 0 7 - 2 2 0 .
Infectious haematopoietic necrosis
4.
41
H s u Y . L . , M A R K ENGELKING H. & L E O N G J. ( 1 9 8 6 ) .
- Occurrence of
different types of infectious hematopoietic necrosis virus in fish. Appl.
Environ. Microbiol., 52, 1 3 5 3 - 1 3 6 1 .
5.
RISTOW S.S. & ARNZEN J.M. ( 1 9 8 9 ) .
- Development
of
monoclonal
antibodies that recognize a type 2 specific and a common epitope on the
nucleoprotein of infectious hematopoietic necrosis virus. J. Aquat. Anim.
Health, 1, 1 1 9 - 1 2 5 .
6.
PVJSTOW S.S. & A R N Z E N D E A V I L A J.M. ( 1 9 9 1 ) . - Monoclonal antibodies to
the glucoprotein and nucleoprotein of infectious hematopoietic necrosis
virus (IHNV) reveal differences among isolates of the virus by
immunofluorescence neutralization and electrophoresis. Dis. Aquat. Org.
11, 1 0 5 - 1 1 5 .
7.
8.
W O L F K. ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 p.
WUNNER W . H . & PETERS D. ( 1 9 9 1 ) . - Family Rhabdoviridae. In: Francki
R.I., Fauque C M . , Knudson D . L . , Brown F. Classification and
nomenclature of viruses. Archives of Virology, Supplementum 2 . Springer,
New York, Vienna, 2 5 0 - 2 6 2 .
Oncorhynchus
masou virus disease
43
CHAPTER
ONCORHYNCHUS
4
MASOUVIRUS
DISEASE
(B406)
SUMMARY
Oncorhynchus masou virus disease (OMVD) is an
oncogenic
condition among salmonid fish in Japan and probably of the coastal
rivers of Eastern Asia which harbour Pacific salmon. The causative
virus (OMV) is also known as Yamame tumor virus (YTV), or Nerka
virus Towada Lake, Akita and Amori prefecture (NeVTA). For a
recent and more detailed review of the condition, see Wolf (5).
The susceptible fish species are kokanee salmon (Oncorhynchus
nerka), masou salmon (O. masou), chum salmon (O. keta), coho
salmon (O. kisutcfy and rainbow trout (O. mykissj.
Clinically, the initial infection by OMV appears as a systemic and
frequently lethal infection which occurs in a context of oedema and
haemorrhages.
Virus multiplication
in endothelial cells of blood
capillaries, haematopoietic
tissue and hepatocytes
underlies the
clinical signs. Four months after this first clinical condition, a
varying number of surviving fish exhibit epithelioma
occurring
mainly around the mouth (upper and lower jaw), and to a lesser
extent, on the caudal fin operculum and body surface.
These
neoplasia may persist for up to one year
post-infection.
Following the septicaemia phase of OMV infection, an immune
response takes place and results in the synthesis of neutralising
antibodies to OMV. A carrier state frequently occurs which leads to
virus shedding via the sexual products at times of spawning.
On the basis of antigenic studies conducted with
neutralising
polyclonal rabbit antisera, OMJ differs from the herpesvirus
of
Salmonidae type 1 which is present in the western USA and only
weakly pathogenic.
/
The reservoirs of OMV are clinically infected fish and asymptomatic
carriers among groups of cultured, feral or wild fish.
Infectious
virus is shed via faeces, urine, sexual products and probably skin
mucus, whereas the kidney, spleen, liver and tumors are the sites in
which virus is the most abundant during the course of overt
infection. The transmission of OMV is horizontal and possibly
vertical or rather "egg-associated". Horizontal transmission may be
direct or vectorial, water being the major abiotic factor.
Animate
vectors and fomites also act in OMV transmission.
Egg-associated
transmission,
even
if infrequent,
is the only
mechanism
44
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
substantiating the onset of OMV disease in alevins originating from
disinfected eggs which had been incubated and hatched in virus-free
water.
Salmonids are the only fish species susceptible to OMV infection.
The grouping of the fish species from the most to the least
susceptible is kokanee, masou salmon, chum salmon, coho salmon
and rainbow trout. The age of the fish is critical and one month-old
alevins offer the most susceptible targets for the virus infection. The
main environmental factor favouring OMV infection is low water
temperature (below 14°C).
The diagnostic procedures for OMV are based upon direct isolation
of the virus in cell culture followed by immunological
identification
by neutralisation or
immunofluorescence.
Control methods currently rely on the implementation of avoidance
and hygiene practices in the operating of salmonid husbandry. The
thorough disinfection of fertilised eggs and the incubation of these
eggs and rearing of fry and alevins, in premises
completely
separated from those harbouring virus carriers and free from
contact with fomites, are key measures to decrease contamination of
OMV in a defined fish production site.
DIAGNOSTIC PROCEDURES
The diagnosis of Oncorhynchus masou virus disease is based upon direct
methods which are either the isolation of the virus (OMV) in cell culture
followed by its immunological identification (conventional approach), or the
immunological demonstration of OMV antigen (Ag) in infected fish tissues.
Due to insufficient knowledge of the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been recognised as a
valuable diagnostic method for assessing the virus status of fish populations.
However, the validation of some serological techniques for diagnosis of certain
fish virus infections could arise in the near future, rendering the use of fish
serology more widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
during overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
kidney, spleen and encephalon.
-
during dormant infection (detection of asymptomatic virus carrier fish):
kidney, spleen and encephalon (any size fish) and/or ovarian fluid from
broodfish at time of spawning.
Oncorhynchus
masou virus disease
45
Sampling procedures: see Chapter 1 Part B.
1.
STANDARD MONITORING M E T H O D S FOR OMVD
1.1. Isolation of OMV in cell culture
Cell line(s) to be used: CHSE-214.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10 organ
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-old CHSE-214 cell
monolayers. Inoculate at least 2 c m of cell monolayer with
100 pi of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 10-15°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 10-15°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, during 14 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation phase. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the tested homogenate
supernatants, identification procedures have to be undertaken
immediately (see below). If a fish health
surveillance/control
programme is being implemented, provisions have to be taken
to suspend the approved health status of the production unit
and/or the zone (if it was approved previously) from which the
virus positive sample originated. The suspension of approved
status will be maintained until it is demonstrated that the virus
in question is not OMV.
iv)
If no CPE occurs, except in positive control cell cultures,
subcultivation steps have to be made even after 7 days of
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
46
incubation, in certain of the infected cell cultures. However, if
no CPE is observed even in positive controls, another series of
virological examinations have to be undertaken, using
susceptible cells and new batches of organ samples.
c) Subcultivation
procedures
i)
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
ii)
Centrifuge at 2,000 x g for
supernatant.
iii)
Repeat optional neutralisation test to IPNV if needed, with
dilution of the above supernatant (1:1 to 1:100).
iv)
Inoculate CHSE-214 cell monolayers as described
(l.l.a).
v)
Incubate and monitor as in 1.1 .b.
15 min at 4°C and collect
above
1.2. Virus identification
a) Neutralisation
test
i)
Collect the culture medium of the cell monolayers exhibiting
CPE and centrifuge it at 2,000 xg for 15 min at 4°C to remove
cell debris, or
ii)
Dilute virus-containing medium from 10- to 10- .
iii)
Mix aliquots (for example 200 ul) of each virus dilution with
equal volumes of an antibody solution specific for OMV, and
similarly treat aliquots of each virus dilution with cell culture
medium.
2
4
The titre of neutralising antibody (NAb) solution must be
around 2,000 in the 5 0 % plaque reduction assay.
iv)
v)
In parallel, other neutralisation tests must be
against:
performed
-
a homologous virus strain (positive neutralisation test)
-
a heterologous virus strain (negative neutralisation test).
Incubate all the mixtures at 15°C for 1 hour.
Oncorhynchus
vi)
47
masou virus disease
After incubation, transfer aliquots of each of the above
mixtures onto CHSE-214 cell monolayers (inoculate 2 cell
cultures per dilution) and allow adsorption to occur for 0.5 to
1 hour at 15°C. Twenty-four or 12 well cell culture plates are
suitable for this purpose, using a 50 ul inoculum.
vii) When adsorption is completed, add cell culture medium
supplemented with 2 % FCS and buffered at pH 7.4-7.6 into
each well and incubate at 10-15°C.
viii) Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in ethanol 20%.
ix)
The tested virus is identified as OMV when CPE is abolished
or noticeably delayed in the cell cultures which had received
the virus suspension treated with the OMV-specific antibody,
whereas CPE is evident in all other cell cultures.
x)
In the absence of any neutralisation by NAb to OMV, it is
mandatory to conduct an indirect fluorescent antibody test
with the suspect sample.
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
2
i)
Prepare monolayers of CHSE-214 cells in 2 c m wells of cell
culture plastic plates or on coverglasses in order to reach
around 8 0 % confluency, which is usually achieved within 4
hours of incubation at 22°C (seed 6 cell monolayers per virus
isolate to be identified, plus 2 for positive and 2 for negative
controls). The FCS content of the cell culture medium can be
reduced to 2-4%. If numerous virus isolates have to be
identified, the use of Terasaki plates is strongly recommended.
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of O M V in a similar way,
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
48
in order to obtain a virus titre of about 5,000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 15°C for 48 hours.
v)
When the incubation time is over, aspirate the cell culture
medium, rinse once with PBS 0.01 M pH 7.2, then 3 times
briefly with cold fixative. This fixative will be acetone (stored
at -20°C) for coverglasses or a mixture of acetone 30%ethanol 7 0 % (vol./vol.), also stored at -20°C.
vi)
Afterwards, let the fixative act for 15 min. A volume of 0.5 ml
is adequate for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to OMV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the cell monolayers by 4 rinsing steps with the
above PBS and eliminate this buffer thoroughly after the last
rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber and do not allow evaporation to
occur. The volume of solution to be used is 0.25 ml/2 c m
well.
2
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Observe the treated cell monolayers on plastic plates
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
xv)
Conduct this observation under incident UV light using a
microscope with xlO eye pieces and x20 to x40 objective lens
having numerical aperture >0.65 and >1.3 respectively.
Oncorhvnchus
masou virus disease
49
Positive and negative controls must be found to give the
expected results prior to any other observation.
c) Enzyme-linked
immunosorbent
assay
(ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for OMV, in PBS 0.01 M pH 7.2 (200 ul/well).
ii)
Incubate overnight at 4°C.
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 ul/well).
v)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vi)
Rinse 4 times with PBST.
vii) Dispense 100 ul/well of a 2 or 4 step dilution of the virus to be
identified and of OMV control virus, and allow to react with
the coated antibody to OMV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
ix)
Add to the wells, biotinylated polyclonal antibody to OMV.
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(O-phenylenediamide, OPD or other approved chromogen).
Stop the course of the test when positive controls react, and
monitor the results.
xv) Alternatively: add H 0 2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2
50
2.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
D I A G N O S T I C P R O C E D U R E S F O R C O N F I R M A T I O N O F O M V D IN S U S P E C T E D
OUTBREAKS
Confirmation of Oncorhynchus
following methods.
2.1. Conventional
identification
virus
masou virus can be achieved by any of the
isolation
with
subsequent
serological
As Section 1.
2.2. Virus isolation with simultaneous identification
a) Sampling
procedures
As B. 1.1. in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ homogenates
B.3.3. for treatment to neutralise birnaviruses (if required).
c) Virus identification
by neutralisation
test
i)
Dilute the anti-IPN-treated organ homogenates 1:100, 1:1,000
and 1:10,000 in cell culture medium.
ii)
Mix with equal volume of a solution of antibody to OMV as in
1.2.a, inoculate the cell monolayers, incubate at 10-15°C and
monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
U.c.
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom of
the wells of a plastic cell culture plate.
Oncorhvnchus
51
masou virus disease
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required
for virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in
1.2.b. points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and block
with 5 % skim milk or 1% bovine serum albumin (BSA), in
PBST for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to O M V and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Peveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in
1.1.
2.4.
ELISA
a) Microplate
processing
As 1.2.c of this chapter up to point iv (inclusive).
b) Sampling
procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection offish specimens
B.2.1. & 2.2. for the selection of materials sampled.
c) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus
homogenates.
extraction
and
obtaining
of
organ
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
52
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol./vol.) as 1.2.C point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.c.
REFERENCES
1.
HEDRICKR.P., MCDOWELLT.S., EATON W . , KIMURAR. & S A N O T . ( 1 9 8 7 ) .
- Serological relationships of five herpesvirus isolated from salmonid
fishes. J. Ichtyol., 8 7 - 9 2 .
2.
K I M U R A T . , YOSHIMISU M., TANAKA M. & SANNOHE H. ( 1 9 8 1 ) . - Studies
on a new virus (OMV) from Oncorhynchus
pathogenicity. Fish Pathol., 15, 1 4 3 - 1 4 7 .
3.
LANNAN C.N., W I N T O N J . R . , F R Y E R J . L . ( 1 9 8 4 ) . - New cell lines. Fish
cells: Establishment and characterization
salmonids. In Vitro, 20, 6 7 1 - 6 7 6 .
4.
masou I. Characteristics and
of nine
cell
lines
from
ROIZMAN B. ( 1 9 9 1 ) . - Family Herpesviridae. In: Francki R . I . , Fauque
C M . , Knudson D . L . , Brown F. Classification and nomenclature of viruses.
Archives of Virology, Supplementum 2 . Springer, New York, Vienna,
103-110.
5.
W O L F K. ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 pp.
Spring viraemia of carp
53
CHAPTER
5
SPRING VIRAEMIA OF CARP
(B404)
SUMMARY
Spring viraemia of carp (SVC) is a rhabdovirus infection of several
carp species and of some other cyprinid fish species.
Overt
infections have been recognised in common carp (Cyprinus carpió),
grass
carp
(Ctenopharyngodon
idellus),
silver
carp
(Hypophthalmichthys molitrixj, bighead carp (Aristichthys nobilisj,
crucian carp fCarassius carassiusj, goldfish (C. auratus), tench
(Tinea tincaj and sheatfish (Silurus glanisj. The geographic range of
SVC is currently limited to countries of the European
continent
which experience low water temperatures during winter. For a
recent and more detailed review of the condition, see Wolf (4).
Infection by SVC virus (SVCV) can be lethal, due to, as in other
rhabdoviroses of fish, the impairment of the salt-water
balance,
which occurs in a clinical context of oedema and
haemorrhages.
Virus multiplication,
especially
in endothelial
cells of blood
capillaries, haematopoietic tissue and nephron cells, underlies the
clinical signs.
Overcoming SVCV infection results in a strong protective
immunity
associated
with the presence
of circulating
antibodies
(Ab)
detectable
by methods
such as virus neutralisation
(VN),
immunofluorescence
(IF) or ELISA. This health status also results,
in certain individuals, in an asymptomatic virus carrier state.
On the basis of antigenic studies conducted with rabbit polyclonal
neutralising
antibodies, SVCV was found to present only one
serotype using the VN test, but both IFAT and ELISA tests have
revealed that SVCV shares common antigenic domain(s) with the
pike-fry rhabdovirus. Variations in virulence of virus strains have
been recorded
during both natural
cases of disease
and
experimental
infections.
The reservoirs of SVC virus are clinically infected fish and
asymptomatic virus carriers from either cultured, feral or wild fish.
Virulent virus is shed via faeces, urine, sexual fluids and probably
gill and skin epithelia, whereas kidney, spleen, gill and encephalon
are the organs in which SVCV is most abundant during the course of
overt infection.
The mode of transmission for SVCV is horizontal but an "eggassociated" transmission (usually called "vertical") cannot be ruled
54
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
out. Horizontal transmission may be direct or vectorial, water being
the major abiotic vector. Animate vectors and /omites are also
involved in transmission of SVCV. Among animate vectors, the
parasitic invertebrates Argulus foliaceus (Crustacea, Branchiuraj
and Piscicola piscícola (Annelida, Hirudinea) are able to transfer
SVCV from diseased to healthy fish. Once SVCV is established in
pond stock or pond farm stock, it may be very difficult to eradicate
without destroying all kinds of life on the fish production site.
Apart from the formerly cited cyprinid species susceptible to SVCV
infection, it seems that very young fish from various pond fish
species are susceptible to SVC, regardless of water temperature. The
most striking example is that of the pike (Esox lucius) which can be
easily infected via the water route. There is a high variability in the
degree of susceptibility to SVC among individuals of the same fish
species. Apart from the physiological state of the fish, the role of
which is poorly understood, age appears to be extremely
important:
the younger the fish, the more susceptible to overt infection. It
remains certain that the environmental factor which is critical for
SVC infection is water temperature: overt infection is not often
observed above 15°C.
The diagnostic procedures for SVCV are based upon direct methods.
The most widely used is isolation of the virus in cell culture followed
by immunological identification using VN, immunofluorescence
or
ELISA tests.
The implementation of hygiene measures and control policy rules
are the only control methods currently feasible. Vaccination is only
experimental.
DIAGNOSTIC PROCEDURES
The monitoring for and diagnosis of SVC is based upon direct methods which
are either the isolation of SVC virus (SVCV) in cell culture followed by its
immunological identification (conventional approach), or the immunological
demonstration of SVCV antigen (Ag) in infected fish tissues.
Due to insufficient knowledge on the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been accepted as a routine
diagnostic method for assessing the virus status of fish populations. However,
the validation of some serological techniques for diagnosis of certain fish virus
infections could arise in the near future, rendering the use of fish serology more
widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
d u r i n g overt infection: whole alevin (body length < 4 cm), viscera
55
Spring viraemia of carp
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
kidney, spleen and encephalon.
-
d u r i n g d o r m a n t infection (asymptomatic virus carrier fish): encephalon
(any size fish) and/or ovarian fluid from broodfish at time of spawning.
Sampling procedures: see Chapter 1 Part B.
1.
STANDARD MONITORING METHODS FOR
SVC
1.1. Isolation of SVCV in cell culture
Cell line(s) to be used: EPC.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10 organ
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-old EPC cell
monolayers. Inoculate at least 2 c m of cell monolayer with
100 pi of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 10-15°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 15°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, for 7 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation period. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the homogenate, identification
procedures must be undertaken immediately (see below). If a
fish health surveillance/control
programme is being imple­
mented, steps have to be taken to suspend the approved health
status of the production unit and/or the zone (if it was
approved previously) from which the suspected virus positive
56
ODE Diagnostic Manual for Aquatic Animal Diseases, 1995
sample originated. The suspension of approved status will be
maintained until it is demonstrated that the virus in question
is not SVCV.
iv)
If no CPE occurs after 7 days of incubation (except in positive
control cell cultures), subcultivation of the inoculated cell
cultures must be performed. However, if no CPE is observed in
positive controls, another series of virological examinations
must be undertaken, using susceptible cells and new batches of
organ samples.
c) Subcultivation
procedures
i)
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
ii)
Centrifuge at 2,000 x g for
supernatant.
iii)
If required, repeat neutralisation test to IPNV as previously
described, with dilution of the above supernatant (1:1 to
1:100).
iv)
Inoculate EPC cell monolayers as described above (1.1.a).
v)
Incubate and monitor as in 1.1.b.
vi)
If no CPE occurs the test may be declared negative.
15 min at 4°C and
collect
1.2. Virus identification
a) Neutralisation
test
i)
Collect the culture medium of the cell monolayers exhibiting
CPE and centrifuge it at 2,000 xg for 15 min at 4°C, or filter
through a 450 nm pore membrane to remove cell debris.
ii)
Dilute virus-containing medium 10- and 10- .
iii)
Mix aliquots (for example 200 ul) of each dilution with equal
volumes of an antibody solution for SVCV, and similarly treat
aliquots of each virus dilution with cell culture medium.
2
4
(The titre of neutralising antibody (NAb) solution must be at
least 2,000 for 5 0 % plaque reduction.)
iv)
In parallel, a neutralisation test must be performed against
homologous SVCV (positive neutralisation test).
Spring viraemia of carp
57
v)
If required, do a similar neutralisation test with antibodies to
IPN, to ensure that no IPNV contaminant might have escaped
the first anti-IPNV test.
vi)
Incubate all the mixtures at 20°C for 1 hour.
vii) Transfer aliquots of each of the above mixtures onto EPC cell
monolayers (inoculate 2 cell cultures per dilution) and allow
adsorption to occur for 0.5 to 1 hour at 15-20°C. Twenty-four
or 12 well cell culture plates are suitable for this purpose,
using a 50 ul inoculum.
viii) When adsorption is completed, add cell culture medium
supplemented with 2 % FCS and buffered at pH 7.4-7.6 into
each well and incubate at 15-20°C.
ix)
Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in 2 0 % ethanol.
x)
The tested virus is identified as SVCV when CPE is prevented
or noticeably delayed in the cell cultures which received the
virus suspension treated with the SVCV-specific antibody,
whereas CPE is evident in all other cell cultures.
xi)
In the absence of any neutralisation by NAb to SVCV, it is
mandatory to conduct an indirect fluorescent antibody test
with the suspect sample.
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
i)
2
Prepare monolayers of EPC cells in 2 c m wells of cell culture
plastic plates or on coverglasses in order to reach around 8 0 %
confluency within 24 hours of incubation at 30°C (seed 6 cell
monolayers per virus isolate to be identified, plus 2 for positive
and 2 for negative controls). The FCS content of the cell
culture medium can be reduced to 2-4%. If numerous virus
isolates have to be identified, the use of Terasaki plates is
strongly recommended.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
58
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of SVCV in a similar way,
in order to obtain a virus titre of about 5,000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 20°C for 24 hours.
v)
Remove the cell culture medium, rinse once with PBS 0.01 M
pH 7.2, then 3 times briefly with cold acetone (stored at 20°C) for coverglasses or a mixture of acetone 30%-ethanol
7 0 % (vol./vol.), also at -20°C, for plastic wells.
vi)
Let the fixative act for 15 min. A volume of 0.5 ml is adequate
for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to SVCV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the dried cell monolayers by 4 rinsing steps with
the above PBS-Tween and eliminate this buffer thoroughly
after the last rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber and do not allow evaporation to
occur. The volume of solution to be used is 0.25 ml/2 c m
well.
2
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Examine the treated cell monolayers on plastic plates
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
59
Spring viraemia of carp
xv) Examine under incident UV light using a microscope with xlO
eye pieces and x20 to x40 objective lens having numerical
aperture >0.65 and >1.3 respectively. Positive and negative
controls must be found to give the expected results prior to any
other observation.
c) Enzyme-linked
immunosorbent
assay
(ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for SVCV, in PBS 0.01 M pH 7.2 (200 ul/well).
Ig may be polyclonal or monoclonal Ig originating most often
from rabbit or mouse, respectively.
ii)
Incubate overnight at 4°C.
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 ul/well).
v)
Rinse 4 times with PBST.
vi)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vii) Dispense 100 ul/well of 2 or 4 step dilutions of the virus to be
identified and of SVCV control virus, and allow to react with
the coated antibody to SVCV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
ix)
Add to the wells, biotinylated polyclonal antibody to SVCV; or
MAb to N protein specific for a domain different from the one
of the coating MAb and previously conjugated with biotin.
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times with PBST.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(Ophenylenediamide, OPD or other approved chromogen). Stop
60
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
the course of the test when positive controls react, and read the
results.
xv)
2.
Alternatively: add H2O2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
D I A G N O S T I C P R O C E D U R E S F O R C O N F I R M A T I O N O F SVCV IN S U S P E C T E D
OUTBREAKS
Confirmation of SVCV can be achieved by any of the following methods:
2.1. Conventional
identification
virus
isolation
with
subsequent
serological
As Section 1.
2.2. Virus isolation with simultaneous serological identification
a) Sampling
procedures
As B . l . l . in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ homogenates
B.3.3. for treatment to neutralise birnaviruses (if required).
c) Virus identification
by neutralisation
test
i)
Dilute organ homogenates 1:100, 1:1,000 and 1:10,000 in cell
culture medium.
ii)
Mix with equal volume of a solution of antibody to SVCV as
in 1.2.a, inoculate the cell monolayers, incubate at 15-20°C
and monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
U.c.
61
Spring viraemia of carp
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom of
the wells of a plastic cell culture plate.
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required
for virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in
1.2.b. points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and block
with 5 % skim milk or 1% bovine serum albumin (BSA), in
PBST for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to SVCV and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Reveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in
1.1.
2.4. ELISA
Microplate
processing
As 1.2.c of this chapter up to point iv (inclusive).
Sampling
procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection of fish specimens
B.2.1. & 2.2. for the selection of materials sampled.
Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ
homogenates.
62
O I E Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol./vol.) as 1.2.C point v and 2 m M of phenyl methyl sulfonide
fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.C
REFERENCES
1.
FIJAN N., PETRINEC Z . , SULIMANOVIC D . & ZWILLENBERG L . O . ( 1 9 7 1 ) . -
Isolation of the causative agent from the acute form of infectious dropsy of
carp. Vet. Archiv., Zagreb, 4 1 ( 5 6 ) , 1 2 5 - 1 3 8 .
2.
FIJAN N.,
L.O.,
SULIMANOVIC D . , BEARZOTTI M . , M U Z I N I C D . , ZWILLENBERG
CHILMONCZYK S.,
V A U T H E R O T J . F . & D E KINKELIN P.
(1983).
-
Some properties of the Epithelioma papulosum cyprini (EPC) cell line
from carp (Cyprinus carpió). Ann. Virol., Institut Pasteur, 1 3 4 E , 2 0 7 - 2 2 0 .
3.
VESTERGAARD-JORGENSEN P.E., OLESEN N.J., A H N E W . & LORENTZEN N.
( 1 9 8 9 ) . - SVC and PFR viruses. Serological examination of 2 2 isolates
indicates close relationship between the two rhabdoviruses. In: Ahne W . &
Kurstak E. (eds.). Viruses of lower vertebrates. Springer-Verlag, Berlin,
349-366.
4.
5.
W O L F K. ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 p.
W U N N E R W . H . & PETERS D . ( 1 9 9 1 ) . - Family Rhabdoviridae. In: Francki
R.I., Fauque C M . , Knudson D . L . , Brown F. Classification and
nomenclature of viruses. Archives of Virology, Supplementum 2 . Springer,
New York, Vienna, 2 5 0 - 2 6 2 .
Viral haemorrhagic septicaemia
63
CHAPTER
6
VIRAL HAEMORRHAGIC SEPTICAEMIA
(B401)
SUMMARY
Viral haemorrhagic septicaemia (VHS) is a colchvater
rhabdovirus
infection of rainbow trout (Oncorhynchus mykiss), brown trout
(Salmo truttaA grayling
fThymallus thymallus,», white
fish
fCoregonus sp.j, pike (Esox lucius,) and turbot (Scophthalmus
maximus/ Infections with VHS virus in Pacific salmon, Pacific cod
and Pacific herring have always been associated with genetically
characteristic virus strains, which appear to be of low pathogenicity
to rainbow trout.
It occurs in continental Europe but is mainly a matter of concern
because of its clinical and economic consequences in rainbow trout
farming. For a recent and more detailed review of the condition, see
Wolf (8).
The infection of fish is often lethal, due to the impairment of the
salt-water balance, which occurs in a clinical context of oedema and
haemorrhages.
Virus multiplication
in endothelial cells of blood
capillaries, leucocytes, haematopoietic
tissues and nephron cells,
underlies the clinical signs.
In survivors, VHS virus (VHSV) infection results in strong protective
immunity, synthesis of circulating antibodies to VHSV as well as, in
certain individuals, an asymptomatic carrier state. This carrier state
frequently turns to high virus excretion at times of spawning.
Three neutralising subtypes of VHSV have been recognised using a
panel of polyclonal and monoclonal antibody preparations.
Apart
from the above variation, VHSV seem to share a VHS-group
neutralising epitope, and several non-neutralising
epitopes located
on the viral glycoprotein
(G protein). Variations in virus strain
virulence have been recorded in both natural cases of disease and
infection trials.
The reservoirs
of VHSV are clinically
infected
fish
and
asymptomatic carriers among cultured, feral or wild fish. Virulent
virus is shed in the faeces, urine, and sexual fluids whereas kidney,
spleen, brain and digestive tract are the sites in which virus is the
most abundant. Once VHSV is established in a farm stock and
therefore in the water catchment system, the disease
becomes
endemic because of the virus carrier fish.
64
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Several factors influence susceptibility to VHS. Apart from the six
fish species found to be susceptible
to VHSV under
natural
conditions three others, namely sea bass (Dicentrarchus labraxj, sea
bream (Ghrysophris aurataj and lake trout (Salvelinus namaycush),
were demonstrated to be susceptible to water-borne
experimental
infection. Among each fish species, there is a high degree of
individual variability in susceptibility to VHSV. The age of fish
appears to be highly important - the younger the fish the higher the
susceptibility to overt infection. Similarly, a good overall fish health
condition seems to decrease the susceptibility to overt VHS, while
handling stress and other types of stress frequently cause subclinical
infection to become overt.
An important environmental factor is water temperature.
Clinical
disease may occur between 14-18°C, although new infections are
rarely established above 15°C under natural conditions.
The diagnostic procedures for VHS are based mainly on direct
methods and the classical approach involving virus isolation in cell
culture
followed
by
immunological
virus
identification
(neutralisation,
immunofluorescence,
ELISA), is the most widely
used. However, more rapid diagnostic methods evidencing
viral
antigen in infected organ imprints or homogenates
(fluorescence,
ELISA) may be suitable for fish with overt disease. Fish serology
(neutralisation, ELISA) could be of high interest for detecting the
carrier state among fish stocks, but still has to be validated.
Control methods for VHS currently lie in official health surveillance
schemes coupled with control policy measures and a genetic
approach (selection and intergeneric hybridisation). Vaccination is
only at an experimental stage at present.
DIAGNOSTIC PROCEDURES
The monitoring for and diagnosis of VHS is based upon direct methods which
are either the isolation of VHS virus (VHSV) in cell culture followed by its
immunological identification (conventional approach), or the immunological
demonstration of VHSV antigen (Ag) in infected fish tissues.
Due to insufficient knowledge on the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been accepted as a routine
diagnostic method for assessing the virus status of fish populations. However,
the validation of some serological techniques for diagnosis of certain fish virus
infections could arise in the near future, rendering the use of fish serology more
widely acceptable for diagnostic purposes.
65
Viral haemorrhagic septicaemia
Infected fish material suitable for virological examination is:
-
during overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
kidney, spleen and encephalon.
-
d u r i n g d o r m a n t infection (asymptomatic virus carrier fish): encephalon
(any size fish) and/or ovarian fluid from broodfish at time of spawning.
Sampling procedures: see Chapter 1 P a r t B .
1.
STANDARD MONITORING M E T H O D S FOR VHSV
1.1. Isolation of V H S V in cell culture
Cell line(s) to be used: BF-2 or EPC.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10 organ
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-old BF-2 or EPC
cell monolayers. Inoculate at least 2 c m of cell monolayer
with 100 pi of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 10-15°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 15°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, for 7 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation period. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the homogenate, identification
procedures must be undertaken immediately (see below). If a
fish health surveillance/control
programme is being imple-
66
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
merited, steps have to be taken to suspend the approved
status of the production unit and/or the zone (if
approved previously) from which the suspected virus
sample originated. The suspension of approved status
maintained until it is demonstrated that the virus in
is not VHSV.
iv)
health
it was
positive
will be
question
If no CPE occurs after 7 days of incubation (except in positive
control cell cultures), subcultivation of the inoculated cell
cultures must be performed. However, if no CPE is observed in
positive controls, another series of virological examinations
must be undertaken, using susceptible cells and new batches of
organ samples.
c) Subcultivation
procedures
i)
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
ii)
Centrifuge at 2,000 x g for
supernatant.
iii)
If required, repeat neutralisation test to IPNV as previously
described, with dilution of the above supernatant (1:1 to
1:100).
iv)
Inoculate BF-2 or EPC cell monolayers as described above
(l.l.a).
v)
Incubate and monitor as in 1. l.b.
vi)
If no CPE occurs the test may be declared negative.
15 min at 4°C and
collect
1.2. Virus identification
a) Neutralisation
test
i)
Collect the culture medium of the cell monolayers exhibiting
CPE and centrifuge it at 2,000 xg for 15 min at 4°C, or filter
through a 450 nm pore membrane to remove cell debris.
ii)
Dilute virus-containing medium 10- and 10- .
iii)
Mix aliquots (for example 200 ul) of each dilution with equal
volumes of an antibody solution for VHSV, and similarly treat
aliquots of each virus dilution with cell culture medium.
2
4
67
Viral haemorrhagic septicaemia
(The titre of neutralising antibody (NAb) solution must be at
least 2,000 for 5 0 % plaque reduction.)
iv)
In parallel, a neutralisation test must be performed against
homologous VHSV (positive neutralisation test).
v)
If required, do a similar neutralisation test with antibodies to
IPNV, to ensure that no IPNV contaminant might have
escaped the first anti-IPNV test.
vi)
Incubate all the mixtures at 15°C for 1 hour.
vii) Transfer aliquots of each of the above mixtures onto BF-2 or
EPC cell monolayers (inoculate 2 cell cultures per dilution)
and allow adsorption to occur for 0.5 to 1 hour at 15°C.
Twenty-four or 12 well cell culture plates are suitable for this
purpose, using a 50 pi inoculum.
viii) When adsorption is completed, add cell culture medium
supplemented with 2 % FCS and buffered at pH 7.4-7.6 into
each well and incubate at I5°C.
ix)
Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in 2 0 % ethanol.
x)
The tested virus is identified as VHSV when CPE is prevented
or noticeably delayed in the cell cultures which received the
virus suspension treated with the VHSV-specific antibody,
whereas CPE is evident in all other cell cultures.
xi)
In the absence of any neutralisation by NAb to VHSV, it is
mandatory to conduct an indirect fluorescent antibody test
with the suspect sample. Indeed some cases of antigenic drift
of surface antigen have been observed, resulting in occasional
failure of the neutralisation test using NAb to VHSV.
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
68
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
2
i)
Prepare monolayers of BF-2 or EPC cells in 2 c m wells of
cell culture plastic plates or on coverglasses in order to reach
around 8 0 % confluency within 24 hours of incubation at 22°C
(seed 6 cell monolayers per virus isolate to be identified, plus 2
for positive and 2 for negative controls). The FCS content of
the cell culture medium can be reduced to 2-4%. If numerous
virus isolates have to be identified, the use of Terasaki plates is
strongly recommended.
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of VHSV in a similar way,
in order to obtain a virus titre of about 5.000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 15°C for 24 hours.
v)
Remove the cell culture medium, rinse once with PBS 0.01 M
pH 7.2, then 3 times briefly with cold acetone (stored at 20°C) for coverglasses or a mixture of acetone 30%-ethanol
7 0 % (vol./vol.), also at -20°C, for plastic wells.
vi)
Let the fixative act for 15 min. A volume of 0.5 ml is adequate
for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to VHSV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the dried cell monolayers by 4 rinsing steps with
the above PBS-Tween and eliminate this buffer thoroughly
after the last rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber and do not allow evaporation to
occur. The volume of solution to be used is 0.25 ml/2 c m
well.
2
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
Viral haemorrhagic septicaemia
69
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Examine the treated cell monolayers on plastic plates
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
xv) Examine under incident UV light using a microscope with xlO
eye pieces and x20 to x40 objective lens having numerical
aperture >0.65 and >1.3 respectively. Positive and negative
controls must be found to give the expected results prior to any
other observation.
c) Enzyme-linked
immunosorbent
assay (ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for VHSV, in PBS 0.01 M pH 7.2 (200
pl/well). Ig may be polyclonal or monoclonal Ig originating
most often from rabbit or mouse, respectively. For the
identification of VHSV, monoclonal antibodies specific for
certain domains of the nucleocapsid protein (N) are suitable.
ii)
Incubate overnight at 4°C.
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 pl/well).
v)
Rinse 4 times with PBST.
vi)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vii) Dispense 100 pl/well of 2 or 4 step dilutions of the virus to be
identified and of VHSV control virus, and allow to react with
the coated antibody to VHSV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
ix)
Add to the wells, biotinylated polyclonal antibody to VHSV; or
MAb to N protein specific for a domain different from the one
of the coating MAb and previously conjugated with biotin.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
70
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times with PBST.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(0phenylenediamide, OPD or other approved chromogen). Stop
the course of the test when positive controls react, and read the
results.
xv) Alternatively: add H2O2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2.
DIAGNOSTIC PROCEDURES
OUTBREAKS
FOR CONFIRMATION
O F VHS
IN
SUSPECTED
Confirmation of VHS can be achieved by any of the following methods:
2.1. Conventional
identification
virus
isolation
with
subsequent
serological
As Section 1.
2.2. Virus isolation with simultaneous serological identification
a) Sampling
procedures
As B. 1.1. in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b) Processing of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B. 3.2. for virus extraction and obtaining of organ homogenates
B.3.3. for treatment to neutralise birnaviruses (if required).
c) Virus identification
i)
by neutralisation
test
Dilute organ homogenates 1:100, 1:1,000 and 1:10,000 in cell
culture medium.
Viral haemorrhagic septicaemia
71
ii)
Mix with equal volume of a solution of antibody to VHSV as
in 1.2.a, inoculate the cell monolayers, incubate at 15°C and
monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
l.l.c.
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom
of the wells of a plastic cell culture plate.
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required
for virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in
1.2.b. points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and
block with 5 % skim milk or 1% bovine serum albumin (BSA),
in PBST for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to VHSV and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Reveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in
1.1.
2.4. ELISA
a) Microplate
processing
As 1.2.c of this chapter up to point iv (inclusive).
b) Sampling
procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection of fish specimens
B.2.1. & 2.2. for the selection of materials sampled.
O I E Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
72
c) Processing
of organ
samples
See the following sections in Chapter 1 (General Information):
B . 3 . 1 . for transportation
B . 3 . 2 . for virus
homogenates.
extraction
and obtaining
of
organ
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol./vol.) as 1.2.c point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.c.
REFERENCES
1.
JENSEN M.H. ( 1 9 6 5 ) . - Research on the virus of Egtved disease. Ann. N.Y.
Acad. Sci., 1 2 6 , 4 2 2 - 4 2 6 .
2.
LANNAN C.N., W I N T O N J . R . , FRYER J.L. ( 1 9 8 4 ) . - New cell lines. Fish
cells: Establishment and characterization
salmonids. In Vitro, 2 0 , 6 7 1 - 6 7 6 .
3.
of nine
cell
lines
from
LORENZEN N., OLESEN N.J. & VESTERGAARD-JORGENSEN P.E. ( 1 9 8 8 ) . -
Production and characterization of monoclonal antibodies to four Egtved
virus structural proteins. Dis. Aquat. Org., 4, 3 5 - 4 2 .
4.
M O U R T O N C , B E A Z O T T I - L A BARRE M., PIECHACZYK M., PADUCEI F., P A U
B . , BASTIDE J.M. & D E KINKELIN P. ( 1 9 9 0 ) . - Antigen capture ELISA for
viral haemorrhagic septicaemia virus serotype 1. J. Virol. Methods, 2 9 ,
325-334.
5.
M O U R T O N C , ROMESTAND., D E KINKELIN P., JEFFROY J . , LEGOUVELLO R .
& P A U B . ( 1 9 9 2 ) . - A highly sensitive immunoassay for the direct
diagnosis of viral haemorrhagic septicaemia, using anti-nucleocpasid
monoclonal antibodies. J. Clin. Microbiol., 3 0 , 2 3 3 8 - 2 3 4 5 .
6.
OLESEN N.J. & VESTERGAARD-JORGENSEN P.E. ( 1 9 9 2 ) .
-
Comparative
susceptibility of three fish cell lines to Egtved virus, the virus of viral
haemorrhagic septicaemia. Dis. Aquat. Org., 1 2 , 2 3 5 - 2 3 7 .
Viral haemorrhagic septicaemia
7.
73
W O L F K . , G R A V E L L M . & M A L S B E R G E R R . G . ( 1 9 6 6 ) . - Lymphocystis virus:
isolation and propagation in centrarchid fish cell line. Science, 151, 1 0 0 4 1005.
8.
9.
W O L F K. ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 pp.
W U N N E R W . H . & PETERS D. ( 1 9 9 1 ) . - Family Rhabdoviridae. In: Francki
R.I., Fauque C M . , Knudson D.L., Brown F. Classification and
nomenclature of viruses. Archives of Virology, Supplementum 2 . Springer,
New York, Vienna, 2 5 0 - 2 6 2 .
Channel catfish virus disease
75
DISEASES OF FISH
OTHER SIGNIFICANT DISEASES
CHAPTER
7
CHANNEL CATFISH VIRUS DISEASE
(B411)
SUMMARY
Channel catfish virus disease (CCVD) is caused by a herpesvirus
and affects channel catfish (lctalurus punctatusj in the USA. For a
recent and more detailed review of the condition, see Wolf (5).
CCVD is of importance because of its clinical
consequences
in channel catfish farming. CCVD,
mostly the juveniles,
is often lethal, due to the
osmotic balance, which occurs in a clinical context
haemorrhages.
Virus multiplication
in endothelial
capillaries, haematopoietic
tissue and hepatocytes
clinical signs.
and
economic
which affects
impairment
of
of oedema and
cells of blood
underlies the
In survivors, CCV infection results in a strong protective
immunity,
synthesis of circulating antibodies to the virus, and, in certain
individuals, an asymptomatic virus carrier state. This carrier state
frequently leads to virus shedding via the sexual products at times of
spawning.
On the basis of antigenic studies conducted with polyclonal
rabbit
antibodies, CCV isolates appear to form a homogeneous group by
neutralisation, fluorescence and ELISA tests. Some variation in the
virulence of CCV strains has been recorded during
natural
outbreaks of disease and demonstrated
experimentally.
The reservoirs of CCV are clinically infected fish and asymptomatic
carriers. Infectious CCV is shed via faeces, urine and the sexual
products whereas kidney, spleen, intestine and encephalon are the
sites in which virus is the most abundant during the course of overt
infection. The transmission of CCV is horizontal and possibly
vertical or, rather, "egg-associated". Horizontal transmission
may
be direct or vectorial, water being the major abiotic vector. Animate
vectors and fomites also act in CCV transmission.
Egg-associated
transmission seems to be an infrequent event but is the only route
substantiating
the onset of CCVD in alevins originating
from
thoroughly disinfected eggs which had been incubated and hatched
in virus-free water. Once CCVD is established in a fish stock, it
becomes endemic because of carrier fish.
76
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Channel catfish is the only fish species susceptible to CCV and
variations in susceptibility to CCV have been recorded
depending
on fish strains. The age of fish is extremely important for overt
infection: the younger the fish, the more susceptible to CCVD.
Water temperature is the critical environmental factor: the mortality
rate is high at 25°-30°C but readily decreases and ceases below
18°C.
The diagnostic procedures for CCV are all based upon direct
methods. The conventional approach is the most widely used and
involves isolation of the virus in cell culture followed
by
immunological identification by neutralisation,
immunofluorescence
or ELISA tests. Rapid techniques by immunofluorescence
or ELISA
test are suitable mainly for diagnosis in clinically infected fish, but
it is likely that ELISAs using certain monoclonal antibodies specific
for the virus nucleocapsid antigen will become a
recommended
method for virus carrier screening, by detection of the viral antigen
in the encephalon.
Control methods currently rely on the implementation
of control
policy rules and of hygiene practices in the operating of catfish
husbandry. The thorough disinfection of fertilised eggs and the
incubation of eggs and further rearing of fry and alevins in premises
completely separated from those harbouring possible virus carriers
and free from possible contact with fomites,
are critical for
preventing the occurrence of CCV in a defined fish production site.
Vaccination, although experimentally
promising, is not used in
practice.
DIAGNOSTIC PROCEDURES
The diagnosis of CCVD is based upon direct methods which are either the
isolation of CC virus (CCV) in cell culture followed by its immunological
identification (conventional approach), or the immunological demonstration of
CCV antigen (Ag) in infected fish tissues.
Due to insufficient knowledge of the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been recognised as a
valuable diagnostic method for assessing the virus status of fish populations.
However, the validation of some serological techniques for diagnosis of certain
fish virus infections could arise in the near future, rendering the use of fish
serology more widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
d u r i n g overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
Channel catfish virus disease
77
kidney, spleen and encephalon.
-
during dormant infection (detection of asymptomatic virus carrier fish):
encephalon (any size fish) and/or ovarian fluid from broodfish at time of
spawning.
Sampling procedures, see Chapter 1 Part B.
1.
S T A N D A R D M O N I T O R I N G M E T H O D S F O R CCVD
1.1. Isolation of CCV in cell culture
Cell line(s) to be used: CCO.
a) Inoculation
i)
of cell
monolayers
Make two additional tenfold dilutions of the 1:10 organ
homogenate supernatants and transfer an appropriate volume
of each of the three dilutions onto 24-hour-oId CCO cell
monolayers. Inoculate at least 2 c m of cell monolayer with
100 ul of each dilution.
2
ii)
Allow to adsorb for 0.5-1 hour at 25-30°C and, without
withdrawing inoculate, add cell culture medium buffered at pH
7.6 and supplemented with 2 % FCS (1 ml/well for 24 well cell
culture plates) and incubate at 25-30°C.
b) Monitoring
incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, during 14 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation phase. This can be achieved by
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the tested homogenate
supernatants, identification procedures have to be undertaken
immediately (see below). If a fish health
surveillance/control
programme is being implemented, provisions have to be taken
to suspend the approved health status of the production
unit
and/or the zone (if it was approved previously) from which the
virus positive sample originated. The suspension of approved
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
78
status will be maintained
in question is not CCV.
iv)
until it is demonstrated
that the virus
If no CPE occurs, except in positive control cell cultures,
subcultivation steps have to be made even after 7 days of
incubation, in certain of the infected cell cultures. However, if
no CPE is observed even in positive controls, another series of
virological examinations have to be undertaken, using
susceptible cells and new batches of organ samples.
c) Subcultivation
procedures
i)
Collect aliquots of cell culture medium from all monolayers
inoculated with dilutions of each supernatant of organ
homogenates.
ii)
Centrifuge at 2,000 x g for
supernatant.
iii)
Repeat optional neutralisation test to IPNV if needed, with
dilution of the above supernatant (1:1 to 1:100).
iv)
Inoculate CCO cell monolayers as described above (1.l.a).
v)
Incubate and monitor as in 1. l.b.
vi)
Make a second (and last) subcultivation step, if the first one
remains virus-negative.
15 min at 4°C and collect
1.2. Virus identification
a) Neutralisation
test
i)
Collect the culture medium of the cell monolayers exhibiting
CPE and centrifuge it at 2,000 xg for 15 min at 4°C to remove
cell debris, or
ii)
Dilute virus-containing medium from 10-2 to 10-4.
iii)
Mix aliquots (for example 200 pi) of each virus dilution with
equal volumes of an antibody solution specific for CCV, and
similarly treat aliquots of each virus dilution with cell culture
medium.
The titre of neutralising antibody (NAb) solution must be
around 2,000 in the 5 0 % plaque reduction assay.
Channel catfish virus disease
iv)
79
In parallel, other neutralisation tests must be
against:
performed
-
a homologous virus strain (positive neutralisation test)
-
a heterologous virus strain (negative neutralisation test).
v)
Incubate all the mixtures at 25°C for 1 hour.
vi)
Transfer aliquots of each of the above mixtures onto CCO cell
monolayers (inoculate 2 cell cultures per dilution) and allow
adsorption to occur for 0.5 to 1 hour at 25°C. Twenty-four or
12 well cell culture plates are suitable for this purpose, using a
50 ul inoculum.
vii) When adsorption is completed, add cell culture medium
supplemented with 2 % FCS and buffered at pH 7.3-7.6 into
each well and incubate at 25-30°C.
viii) Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in ethanol 20%.
ix)
The tested virus is identified as CCV when CPE is abolished
or noticeably delayed in the cell cultures which had received
the virus suspension treated with the CCV-specific antibody,
whereas CPE is evident in all other cell cultures.
x)
In the absence of any neutralisation by NAb to CCV, it is
mandatory to conduct an indirect fluorescent antibody test
with the suspect sample.
b) Indirect fluorescent
antibody
test
This virus identification test is to be conducted either directly after
virus isolation in cell culture, or as a confirmatory test following the
neutralisation test described above.
i)
2
Prepare monolayers of CCO cells in 2 c m wells of cell culture
plastic plates or on coverglasses in order to reach around 8 0 %
confluency, which is usually achieved within 4 hours of
incubation at 30°C (seed 6 cell monolayers per virus isolate to
be identified, plus 2 for positive and 2 for negative controls).
The FCS content of the cell culture medium can be reduced to
2-4%. If numerous virus isolates have to be identified, the use
80
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
of Terasaki plates is strongly recommended.
ii)
When the cell monolayers are ready for infection, i.e. on the
same day or on the day after seeding, inoculate the virus
suspensions to be identified by making ten-fold dilution steps
directly in the cell culture wells or flasks.
iii)
Dilute the control virus suspension of CCV in a similar way,
in order to obtain a virus titre of about 5,000-10,000 PFU/ml
in the cell culture medium.
iv)
Incubate at 25°C for 18 hours.
v)
When the incubation time is over, aspirate the cell culture
medium, rinse once with PBS 0.01 M pH 7.2, then 3 times
briefly with cold fixative. This fixative will be acetone (stored
at -20°C) for coverglasses or a mixture of acetone 30%ethanol 7 0 % (vol./vol.), also stored at -20°C.
vi)
Afterwards, let the fixative act for 15 min. A volume of 0.5 ml
is adequate for 2 c m of cell monolayer.
2
vii) Allow the cell monolayers to air dry for at least 30 min and
process immediately or freeze at -20°C.
viii) Prepare a solution of purified antibody or serum to CCV in
PBS 0.01 M, pH 7.2 containing 0.05% Tween 80, at the
appropriate dilution (which has been established previously or
is given by the reagent supplier).
ix)
Réhydrate the dried cell monolayers by 4 rinsing steps with
the above PBS and eliminate this buffer thoroughly after the
last rinsing.
x)
Treat the cell monolayers with the antibody solution for 1 hour
at 37°C in a moist chamber. The volume of solution to be used
is 0.25 ml/2 c m well.
2
xi)
Rinse 4 times with PBS-Tween as above.
xii) Treat the cell monolayers for 1 hour at 37°C with a solution of
fluorescein isothiocyanate-conjugated (FITC) antibody to the
immunoglobulin used in the first layer and prepared according
to the instructions of the supplier. These FITC antibodies are
most often rabbit or goat antibodies.
xiii) Rinse 4 times with PBS-Tween.
xiv) Observe
the
treated
cell
monolayers
on
plastic
plates
Channel catfish virus disease
81
immediately, or mount the coverglasses using glycerol saline
at pH 8.5 prior to microscopic observation.
xv)
Conduct this observation under incident UV light using a
microscope with xlO eye pieces and x20 to x40 objective lens
having numerical aperture >0.65 and >1.3 respectively.
Positive and negative controls must be found to give the
expected results prior to any other observation.
c) Enzyme-linked
immunosorbent
assay
(ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for CCV, in PBS 0.01 M pH 7.2 (200 ul/well).
Ig may be polyclonal or monoclonal Ig originating most often
from rabbit or mouse, respectively.
ii)
Incubate overnight at 4°C.
iii)
Rinse 4 times with PBS 0.01 M containing 0.05 % Tween 20
(PBST).
iv)
Block with skim milk ( 5 % in PBST) or other blocking solution
for 1 hour at 37°C (200 ul/well).
v)
Add 2 % of Triton X 100 to the virus suspension to be
identified.
vi)
Rinse 4 times with PBST.
vii) Dispense 100 ul/well of a 2 or 4 step dilution of the virus to be
identified and of CC control virus, and allow to react with the
coated antibody to CCV for 1 hour at 20°C.
viii) Rinse 4 times with PBST.
ix)
Add to the wells, biotinylated polyclonal antibody to CCV.
x)
Incubate 1 hour at 37°C.
xi)
Rinse 4 times with PBST.
xii) Add streptavidin-conjugated horse radish peroxidase to those
wells which have received the biotin-conjugated antibody and
incubate for 1 hour at 20°C.
xiii) Rinse 4 times.
xiv) Add
the
substrate
(H2O2)
and
chromogen
(Ophenylenediamide, OPD or other approved chromogen). Stop
the course of the test when positive controls react, and monitor
the results.
82
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
xv) Alternatively: add H 0 2 + chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2
2.
D I A G N O S T I C P R O C E D U R E S F O R C O N F I R M A T I O N O F CCVD IN S U S P E C T E D
OUTBREAKS
Confirmation of CCV can be achieved by any of the following methods.
2.1. Conventional
virus
isolation
identification as in Section 1.
with
subsequent
serological
2.2. Virus isolation with simultaneous identification
a) Sampling
procedures
As B. 1.1. in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b) Processing
of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ
homogenates
B.3.3. for treatment to neutralise birnaviruses.
c) Virus identification
by neutralisation
test
i)
Dilute the anti-IPN-treated organ homogenates 1:100, 1:1,000
and 1:10,000 in cell culture medium.
ii)
Mix with equal volume of a solution of antibody to CCV as in
1.2.a, inoculate the cell monolayers, incubate at 25-30°C and
monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
U.c.
2.3. Indirect fluorescent antibody test
i)
Bleed the fish thoroughly.
ii)
Make kidney imprints on cleaned glass slides or at the bottom of
the wells of a plastic cell culture plate.
83
Channel catfish virus disease
iii)
Store the kidney pieces (as indicated in B.3.1. in Chapter 1
[General Information]) together with the other organs required
for virus isolation in case this later becomes necessary.
iv)
Allow the imprint to air-dry for 20 min.
v)
Fix with acetone or ethanol-acetone and dry as indicated in
1.2.b. points v-vii.
vi)
Réhydrate the above preparations (see 1.2.b. point ix) and block
with 5 % skim milk or 1% bovine serum albumin (BSA), in
PBST for 30 min at 37°C.
vii) Rinse 4 times with PBST.
viii) Treat the imprints with the solution of antibody to CCV and
rinse as indicated in 1.2.b.
ix)
Block and rinse as formerly.
x)
Reveal the reaction with suitable FITC, rinse and observe as
indicated in 1.2.b. points xii-xv.
If the immunofluorescence test is negative, process the organ
samples stored at 4°C, for virus isolation in cell culture as in
1.1.
2.4. E L I S A
a) Microplate
processing
As 1.2.c of this chapter up to point iv (inclusive).
b) Sampling
procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection of fish specimens
B.2.1. & 2.2. for the selection of materials sampled.
c) Processing
of organ
samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ
homogenates.
d) Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
84
ii)
Treat the remaining part of homogenate with 2 % Triton X 1 0 0
(vol/vol.) as 1.2.c point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.c.
REFERENCES
1.
ARKUSH K.D.,
M C N E I L C.
& HEDRICK R . P .
(1992).
- Production
and
characterization of monoclonal antibodies against channel catfish virus. J.
Aquat. Anim. Health, 4 , 8 1 - 8 9 .
2.
BOWSER P . R & PLUMB J.A. ( 1 9 8 0 ) . - Fish cell lines: establishment of a line
from ovaries of channel catfish. In Vitro. 16, 3 6 5 - 3 6 8 .
3.
ROIZMAN B. ( 1 9 9 1 ) . - Family Herpesviridae. In: Francki R . I . , Fauque
C M . , Knudson D . L . , Brown F. Classification and nomenclature of viruses.
Archives of Virology, Supplementum 2 . Springer, New York, Vienna,
103-110.
4.
W O L F K. & DARLINGTON R . W . ( 1 9 7 1 ) . - Channel catfish virus: a new
herpesvirus of Ictalurid fish. J. Virol., 8 ( 4 ) , 5 2 5 - 5 3 3 .
5.
W O L F K. ( 1 9 8 8 ) . - Fish viruses and viral diseases. Cornell University
Press, Ithaca, New York, 4 7 6 pp.
Viral encephalopathy
85
CHAPTER
8
VIRAL ENCEPHALOPATHY AND RETINOPATHY
(No O I E number)
SUMMARY
A distinctive
syndrome
of vacuolating
encephalopathy
and
retinopathy, or viral nervous necrosis (VNN), occurs in larval and,
sometimes, juvenile sea bass (Lates calcarifer and Dicentrarchus
labrax), turbot (Scophthalmus maximus), Japanese
parrotfish
(Oplegnathus fasciatus), redspotted grouper (Epinepheles akaara),
and striped jack (Pseudocaranx dentex). Typically there is massive
(often 100%) mortality in affected stocks.
The causative agents are icosahedral, non-enveloped viruses about
25-30 nm in diameter and, until recently, described as picornaviruslike. The agent (SJNNV) of VNN in striped jack has been identified
as a nodavirus and serological relationships
have been shown
between
this
virus
and
the agents
causing
vacuolating
encephalopathy
and retinopathy in all the other species
except
turbot (in this instance no tests have been made). Diagnosis depends
on microscopy
(light and electron) and a range of recently
developed immunological and molecular
procedures.
Control measures are based on improved hatchery hygiene and
reduced stocking rates. In the case of VNN in striped jack, where
vertical transmission
unequivocally
occurs, identification
and
culling of carrier broodfish is desirable.
INTRODUCTION
Vacuolating encephalopathy and retinopathy, or viral nervous necrosis (VNN),
of larval marine fish has been described in Australasian sea bass (Lates
calcarifer), European sea bass (Dicentrarchus labrax), turbot
(Scophthalmus
maximus), Japanese parrotfish (Oplegnathus fasciatus),
redspotted grouper
(Epinepheles akaara), and striped jack (Pseudocaranx
dentex). Except for
turbot, there have been confirmed and unconfirmed reports of these syndromes
from most places where the above species are intensively cultured. Recently,
apparently identical disease outbreaks have been reported in tiger puffer
(Talifugu rubripes), Japanese flounder (Paralichthys olivaceus), kelp grouper
(Epinephelus moara) and rock porgy (Oplegnathus
punctatus).
Virus particles of about 25-30 nm in diameter have been visualised in affected
fish and the agent (SJNNV) associated with the disease (VNN) in striped jack
has been characterised and placed in the family Nodavirdae. Immunological
studies have shown a relationship between SJNNV and the agents of the two sea
bass diseases and VNN of Japanese parrotfish and redspotted grouper.
86
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Clinical signs and macroscopic lesions
There is great commonality of clinical signs with 'mass mortality' and a variety
of neurological abnormalities, as follows:
L. calcarifer
uncoordinated darting, corkscrew swimming, pale colour,
anorexia, wasting
D. labrax
whirling swim pattern, swimbladder hyperinflation,
anorexia
O. fasciatus
spiral swimming, dark colour
E. akaara
whirling swim pattern
P. dentex
abnormal swimming behaviour, swimbladder
hyperinflation
S. maximus
spiral and/or looping swim pattern, belly-up at rest, dark
colour
Apart from colour changes and wasting there are no consistent macroscopic
findings.
Interesting differences in relation to the occurrence and severity of the diseases
are shown in Table 1.
EPIDEMIOLOGY
The incubation period for the L. calcarifer disease is four days whereas the
earliest known occurrence of natural disease is nine days after hatch (Table 1),
suggesting that vertical transmission of virus is unlikely in this species.
In contrast, it has been demonstrated that vertical transmission of the causative
agent occurs in P. dentex and this fact is reflected by early occurrence of clinical
disease.
The mode of transmission of the viruses, other than in gametes, has not been
demonstrated but the possibilities include influent water, juvenile fish held on
the same site, and carriage on utensils, vehicles, etc. It is possible that these
small viruses are quite resistant to environmental conditions and therefore
readily translocated by commercial activities.
CONTROL
Control of
The newly
broodfish.
ameliorate
VNN in striped jack is difficult because of its vertical transmission.
developed PCR method should permit screening of potential carrier
Also, there is some evidence that reduced stress at spawning can
the condition.
Viral encephalopathy
87
Table 1. Important features of viral encephalopathies
of larval and juvenile fish
Earliest
occurrence
of disease
Usual onset
of disease
Latest
occurrence
of new
outbreaks
Usual
mortality
rate
Highest
mortality
rate
L. calcarifer
9 d.p.h.
15-18 d.p.h.
>24 d.p.h.
50-100%/
month
100% in <1
month
D. labrax
10 d.p.h.
25-40 d.p.h.
Bodyweight
10%/month
0. fasciatus 6-25 m m t.l.
<40 m m t.l.
Up to 100%
E. akaara
14 d.p.h.
(7-8 m m
t.l.)
9-10 m m t.l.
<40 m m t.l.
80%
P. dentex
1 d.p.h.
1-4 d.p.h.
<20 d.p.h.
(8 m m t.l.)
100%
S. maximus
<21 d.p.h.
d.p.h. = days post-hatch;
Bodyweight
50-100 mg
Up to 100%
Up to 100%
t.l. = total length
In contrast, control of clinical disease in L. calcarifer has been remarkably
successful. In intensive hatcheries this has consisted of some or all of the
following strategies: no recycling of culture water, chemical disinfection of
influent water and larval tanks between batches, and reduction of larval density
from 15-30 larvae/litre to not more than 15 larvae/litre (preferably less than
10/litre). Extensive culture in 'green ponds' is also associated with a low
prevalence of clinical disease and/or histological lesions.
DIAGNOSTIC PROCEDURES
Presumptive diagnosis can be made on the basis of the light microscopic
appearance of the brain and/or retina. However, individual fish with the
presence of only a few vacuoles in the neuropil pose a difficult diagnostic
problem.
Electron microscopy is a very useful confirmatory technique. In particular,
negative staining yields a rapid result and can be performed on both unfixed
(preferable) and formalin-fixed material.
SJNNV can be detected by the fluorescent antibody technique (FAT), enzymelinked immunosorbent assay (ELISA), or polymerase chain reaction (PCR)
amplification of RNA. The FAT is sufficiently broad in specificity to be used to
detect at least four other viruses in this group.
88
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
To date it has proved impossible to culture these viruses in a range of fish tissue
cell cultures.
1.
HlSTOPATHOLOGY
Normal histological methods, including haematoxylin and eosin staining,
are used. Small larvae are embedded whole and serially sectioned to provide
sections of brain and eyeballs. Larger fish (juvenile) usually require removal
of eyes and brain for embedding after preliminary fixation.
All the diseases are characterised by vacuolation of the brain. Usually there
is also vacuolation of the nuclear layers of the retina, although this lesion is
not included in the description of the disease in Japanese parrotfish or
turbot. In general, younger fish have more severe lesions whereas older fish
have less extensive lesions and these may show a predilection for the retina.
Intracytoplasmic inclusions (up to 5 um in diameter) have been described in
sections of European and Australasian sea bass nervous tissues, and
neuronal necrosis has been described in most species. Vacuolation of the gut
is not caused by these nodaviruses, but is a typical physiological response.
2.
ELECTRON MICROSCOPY
Virus particles can be visualised in affected brain and retina by both positive
and negative staining.
In positively stained material the virus is mainly associated with vacuolated
cells and, especially, any inclusions. The particles vary in size from 22-32
nm (European sea bass) to 34 nm (Japanese parrotfish) arranged
intracytoplasmically in crystalline arrays, or as aggregates and single
particles both intra- and extracellularly. The virus is non-enveloped and
icosahedral in shape.
(i)
Negative staining
Fresh or frozen brain or eye is macerated with 1 or 2 % phosphotungstic
acid (adjusted to pH 6.8 with KOH) and mounted on copper grids for
transmission electron microscopy.
Non-enveloped, round to icosahedral particles about 25-30 nm will be
present. It may be possible to detect capsomeres.
(ii)
Whole larvae or eyes and brains of juveniles are fixed in cold 1-2%
glutaraldehyde in 0.1 M cacodylate buffer (pH 7.1) for 24 hours. The
samples are then rinsed three times with 0.1 M cacodylate buffer and
postfixed in 1% osmium tetroxide for one hour. The tissues are next
dehydrated in a series of acetone and placed in a 1:1 mixture of 100%
acetone: Spurr's resin and allowed to infiltrate overnight. Infiltration is
completed by transfer to Spurr's resin for 12 hours and then the tissues
Viral encephalopathy
89
are embedded. Ultrathin sections are cut and stained with uranyl
acetate and lead citrate and examined under a transmission electron
microscope. Virus particles are especially associated with inclusions
and the cytoplasm of vacuolated cells.
3.
FLUORESCENT ANTIBODY TECHNIQUE
Fish samples fixed in 10% buffered formalin are immersed in 0.1 M PBS
containing 5 % sucrose (pH 7.2) overnight. After successive immersions in
10%, 3 0 % and 4 0 % sucrose-PBS for 1-2 h, samples are embedded in OCT
compound (Miles Inc.), cut at 7 um by a cryostat apparatus, and washed
with cold PBS. Samples are incubated with anti-SJNNV rabbit serum
(dilution 1:100) at 37°C for 30 min, washed with PBS, and then reacted
with FITC labelled anti-rabbit Ig goat antibody at 37°C for 30 min, washed
with PBS, and examined by fluorescence microscopy. Specific fluorescence
is observed in the cytoplasm of the affected cells in brain and retina. This
FAT is also applicable to paraffin embedded sections.
4.
E N Z Y M E - L I N K E D I M M U N O S O R B E N T ASSAY ( E L I S A )
At present, ELISA using anti-SJNNV rabbit serum can be used only to
detect VNN from diseased larvae of striped jack.
The fish sample (usually 10 larvae) is homogenised with nine volumes of
0.05 M carbonate-bicarbonate buffer (pH 9.6) and centrifuged at 10,000 g
for 30 min. Serially two-fold diluted supernatant is added in a 96-well
microplate (0.2 ml/well) and incubated at 25°C for 2 h or at 4°C overnight.
After washings with PBS containing 0.05% Tween 20 and blocking with 2 %
bovine serum albumin, wells are filled with 0.2 ml of anti-SJNNV rabbit
serum (dilution 1:1,000) and incubated at 37°C for 2 h. Following washes,
0.2 ml of alkaline phosphatase conjugated anti-rabbit Ig goat antibody is
added to each well and incubated at 37°C for 2 h. Following the final
washing, 0.2 ml of p-nitrophenyl phosphate disodium salt in diethanolamine
solution (pH 9.8) are added and the plates are incubated for 60 min. The
absorbance of each well is read at 405 nm uisng a microplate reader. The
absorbance for normal larvae is usually lower than 0.1 in every dilution.
5.
POLYMERASE CHAIN REACTION ( P C R ) AMPLIFICATION
SJNNV contains two single-stranded, positive-sense, non polyadenylated
RNAs ÇRNAl and RNA2). RNA2 encodes a 42 kDa structural protein of the
virus.
The
two
primers,
a
reverse
primer
(5'CGAGTC AACACGGGTGAAGA-3 ')
and
a
forward
primer
(5'CGTGTCAGTCATGTGTCGCT-3 '), are used for amplification of a target
sequence (about 430 bases) of SJNNV RNA2 by PCR. This PCR procedure
can be used for detection of the viruses from not only striped jack but also
other fish (O. fasciatus, E. akaara, T. rubripes, P. olivaceus, E. moara, and
O. punctatus).
90
O I E Diagnostic Manual for Aquatic Animal Diseases, 1995
The fish sample (0.1 g) is homogenised with 0.5 ml distilled water treated
with 0 . 1 % diethyl pyrocarbonate and centrifuged at 10,000 g for 10 min.
The resultant supernatant is mixed with 0.04 ml of proteinase K (1 mg/ml)
and 0.04 ml of 1% SDS, and incubated at 37°C for 30 min. After
centrifugation, total nucleic acids are extracted through the phenolchloroform method. The total nucleic acids are preheated at 90°C for 5 min
and incubated at 42°C for 30 min in 20 pi of PCR buffer (10 m M Tris-HCl,
pH 8.3, 50 m M KC1) containing 2.5 U of M-MLV reverse transcriptase
(USB), 1.0 U of ribonuclease inhibitor, 0.5 u M of reverse primer, 1 mM
each of four deoxynucleotide triphosphates (dNTP), and 5 m M of MgCl2.
The mixture is incubated at 99°C for 10 min to inactivate the reverse
transcriptase and then diluted five-fold with PCR buffer containing 0.1 uM
of forward primer, 2.5 U of Tth Version 2.0 DNA polymerase (Toyobo) and
2 m M of MgCl2- The mixture is incubated in an automatic thermal cycler
programmed for one cycle at 72°C for 10 min and 95 °C for 2 min, then 25
cycles at 95°C for 40 s, 55°C for 40 s, and 72°C for 40 s, and finally held at
72°C for 5 min. Amplified DNA (430 bp) is analysed by agarose gel
electrophoresis.
REFERENCES
1.
BREUIL G., BONAMI J.R., PEPIN T . F . & P I C H O T Y . (1991). Viral infection
(picorna-like virus) associated with mass mortalities in hatchery-reared
sea-bass (Dicentrarchus labrax) larvae and juveniles. Aquaculture, 97,
109-116.
2.
MORI
K.,
NAKAJ. T.,
MUROGA
K.,
ARIMOTO
M.,
MUSHIAKE
K.
&
FURUSAWA I. (1992). Properties of a new virus belonging to Nodaviridae
found in larval striped jack (Pseudocaranx dentex) with nervous neurosis.
Virology, 187, 368-371.
3.
M U N D A Y B . L . , L A N G D O N J.D., H Y A T T A. & HUMPHREY J.D. (1992). Mass
mortality associated with a viral-induced vacuolating encephalopathy and
retinopathy of larval and juvenile barramundi, Lates calcarifer Bloch.
Aquaculture, 103, 197-211.
4.
NISHIZAWA T., M O R I K., NAKAJ T., FURUSAWA I. & M U R O G A K. (1994).
Polymerase chain reaction (PCR) amplification of RNA of striped jack
nervous necrosis virus (SJNNV). Dis. Aquatic Organisms, 18, 103-107.
5.
YOSHIKOSHI K. & INOUE K. (1990). Viral nervous necrosis in hatcheryreared larvae and juveniles of Japanese parrotfish, Oplegnathus
fasciatus
(Temminck and Schegel). J. Fish Dis., 13, 69-77.
Infectious pancreatic necrosis
91
CHAPTER
9
INFECTIOUS PANCREATIC NECROSIS
(No OIE number)
SUMMARY
Infectious pancreatic necrosis (IPN) is a highly contagious viral
disease of young fish of salmonid species held under intensive
rearing conditions. The disease most characteristically
occurs in
rainbow trout (Onchorhyncus mykiss), brook trout (Salvelinus
fontinalis), brown trout (Salmo trutta), Atlantic salmon (Salmo
salar), and several Pacific salmon species (Oncorhynchus spp.). IPN
virus, or viruses showing serological relatedness to IPN virus, have
been reported to cause diseases in some farmed marine fish species,
such as cod (Gadus morhua), yellowtail (Serióla quinqueradiataj,
turbot
(Scophthalmus maximus,), and halibut
(Hippoglossus
hippoglossus), and subclinical asymptomatic infections have been
detected in a wide range of estuarine and freshwater fish species in
the families
Anguillidae, Atherinidae, Bothidae, Carangidae,
Cotostomidae, Cichlidae, Clupeidae, Cobitidae, Coregonidae,
Cyprinidae, Esocidae, Moronidae, Paralichthydae,
Percidae,
Poecilidae, Sciaenidae, Soleidae and Thymallidae.
The causative agent, infectious pancreatic necrosis virus (IPNV), is
a bi-segmented double-stranded RNA virus belonging to the family
Birnaviridae.
Monitoring for IPN is based upon isolation of the virus in tissue
culture and its immunological identification. Diagnosis of clinical
cases is normally based on characteristic pathological
changes,
particularly to the pancreas (as detected by standard
histological
techniques) and/or immunological demonstration of IPNV antigen in
infected
tissues,
confirmed
by isolation
and
immunological
identification of IPNV in tissue culture.
Control methods currently rely on the implementation
of control
policy rules and of hygiene practices in the operating of salmonid
husbandry, through the avoidance offertilised eggs originating from
IPNV carrier broodstock, and use of a protected water supply (e.g.
spring, borehole pond) into which ingress of fish,
particularly
possible virus carriers, is prevented. In outbreaks, a reduction of the
population density fthinning out') may help reduce the overall
mortality. No treatment or commercial vaccine is available at
present.
92
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
INTRODUCTION
Infectious pancreatic necrosis (IPN) is a highly contagious viral disease,
principally of young fish of salmonid species, held under intensive rearing
conditions. Susceptibility generally decreases with age, with resistance to
clinical disease in salmonid fish usually being reached at about 1,500 degreedays (value obtained by multiplying the age in days by the average temperature
in degrees Centigrade during the lifespan) except for Atlantic salmon smolts
which can suffer from disease shortly after transfer from fresh water to
seawater. The causative agent is a birnavirus (bi-segmented double-stranded
RNA) displaying wide antigenic diversity and marked differences in degrees of
virulence.
The disease has a wide geographical distribution, occurring in most, if not all,
major salmonid farming countries of North and South America, Europe and
Asia.
The first sign of an outbreak in salmonid fry is frequently a sudden and usually
progressive increase in daily mortalities, particularly in the faster growing
individuals. Clinical signs include darkening pigmentation, a pronounced
distended abdomen and a corkscrewing/spiral swimming motion. Cumulative
mortalities may vary from less than 10% to more than 9 0 % depending on the
combination of several factors such as virus strain, host and environment.
Clinical diagnosis is most often confirmed by histology of the internal organs
(particularly of the pancreas) with isolation of the virus in tissue culture,
followed by its identification using a serological method such as serum
neutralisation, ELISA or fluorescent antibody tests (FAT). Virus isolation in
tissue culture is the standard method for detection of asymptomatic carriers.
The disease is transmitted both horizontally through the water route and
vertically via the egg. Surface disinfection of eggs is not entirely effective in
preventing vertical transmission.
There is no treatment or commercial vaccine yet available. Prevention can be
achieved by avoidance of fertilised eggs originating from IPN virus carrier
broodstock and use of a protected water supply (e.g. spring, borehole) into
which ingress of fish is prevented. In outbreaks, a reduction of the population
density ('thinning out') may help reduce the overall mortality.
DIAGNOSTIC PROCEDURES
Monitoring for IPN is based upon isolation of IPN virus (IPNV) in cell culture
followed by its immunological identification. Diagnosis of clincal cases is
normally based on histology and/or immunological demonstration of IPNV
antigen (Ag) in infected tissues (confirmed by isolation and immunological
identification of IPNV in tissue culture).
Infectious pancreatic necrosis
93
Due to insufficient knowledge on the fish serology of virus infections, the
detection of fish antibodies to viruses has not thus far been accepted as a routine
diagnostic method for assessing the virus status of fish populations. However,
the validation of some serological techniques for diagnosis of certain fish virus
infections could arise in the near future, rendering the use of fish serology more
widely acceptable for diagnostic purposes.
Infected fish material suitable for virological examination is:
-
during overt infection: whole alevin (body length < 4 cm), viscera
including kidney (4 cm < body length < 6 cm) or, for larger size fish,
liver, kidney and spleen.
-
during dormant infection (asymptomatic virus carrier fish): liver,
kidney and spleen (any size fish) and/or ovarian fluid from broodfish at
time of spawning.
Sampling procedures: see Chapter 1 Part B.
1.
S T A N D A R D M O N I T O R I N G M E T H O D S F O R D?N
1.1. Isolation of D?NV in cell culture
Cell line(s) to be used: BF-2 and CHSE-214.
a)
b)
Inoculation of cell monolayers
i)
Make a 1:100 dilution of the organ homogenate supernatant
(1:100 of tissue). Incubate at 15-20°C for 1 hour or overnight
at 4°C. Inoculate an appropriate volume of the primary
dilution and a 1:10 dilution thereof onto 24- to 48-hour-old
BF-2 monolayers.
ii)
Allow to adsorb for 1 hour at 15-20°C, then remove the 1:100
dilution and replace with cell culture medium. Incubate at
15°C. All dilutions are to be made in cell culture medium
buffered at pH 7.6 and supplemented with 10% FCS,
antibiotics and antimycotics.
Monitoring incubation
i)
Follow the course of infection in positive controls and other
inoculated cell cultures, by daily microscopic examination at
magnification 40x to lOOx, for 7 days. The use of a phase
contrast microscope is recommended.
ii)
Maintain the pH of the cell culture medium between 7.3 and
7.6 over the whole incubation period. This can be achieved by
94
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
addition to the inoculated cell culture medium of sterile
bicarbonate buffer (for tightly closed cell culture flasks) or 2 M
Tris buffer solution (for cell culture plates) or, even better, by
using HEPES-buffered media.
c)
iii)
If a cytopathic effect (CPE) appears in those cell cultures
inoculated with the dilutions of the homogenate, identification
procedures must be undertaken immediately (see below).
iv)
If no CPE occurs after 7 days of incubation (except in positive
control cell cultures), subcultivation of the inoculated cell
cultures must be performed.
Subcultivation procedures
i)
Collect cell culture monolayers and subject them to one freezethaw cycle. Pool aliquots of the supernatants from all cell
monolayers inoculated with dilutions of organ homogenates.
ii)
Dilute 1:20 and 1:100 and inoculate BF-2 cell monolayers as
described above (1.1.a.).
iii)
Incubate and monitor as in 1.1 .b.
iv)
If no CPE occurs the test may be declared negative.
1.2. Virus identification
a)
Neutralisation test
2
4
i)
Dilute virus-containing medium 10- and IO- .
ii)
Mix aliquots (for example 200 ul) of each dilution with equal
volumes of an antibody solution for IPNV, and similarly treat
aliquots of each virus dilution with cell culture medium.
(The titre of neutralising antibody (NAb) solution must be at
least 2,000 for 5 0 % plaque reduction.)
iii)
Incubate all the mixtures at 15°C for 1 hour.
iv)
Transfer aliquots of each of the above mixtures onto cell
monolayers (inoculate 2 cell cultures per dilution).
v)
Incubate at 15°C.
vi)
Check the cell cultures for the onset of CPE and read the
results as soon as it occurs in non-neutralised controls (cell
Infectious pancreatic necrosis
95
monolayers being protected in positive neutralisation
controls). Results are recorded either after a simple
microscopic examination (phase contrast preferable) or after
discarding cell culture medium and staining cell monolayers
with a solution of 1% crystal violet in 2 0 % ethanol.
vii) The tested virus is identified as IPNV when CPE is prevented
or noticeably delayed in the cell cultures which received the
virus suspension treated with the IPNV-specific antibody,
whereas CPE is evident in all other cell cultures.
b)
Enzyme-linked immunosorbent assay (ELISA)
i)
Coat the wells of microplates designed for ELISA tests, with
appropriate dilutions of purified immunoglobulins (Ig) or
serum specific for IPNV, in carbonate buffer 0.02 M, pH 9.5
(200 pl/well). Ig may be polyclonal or monoclonal Ig
originating most often from rabbit or mouse, respectively. For
the identification of IPNV, monoclonal antibodies specific for
certain domains of the nucleocapsid protein (N) are suitable.
ii)
Incubate overnight at 4°C.
iii)
Rinse 2 times with PBS 0.01 M.
iv)
Block with bovine serum albumin (1.5% in carbonate buffer)
or other blocking solution for 1 hour at 37°C (200 pl/well).
v)
Rinse 4 times with PBST containing 0.05 % Tween 20
(PBST).
vi)
Add an equal volume of PBST to the virus suspension to be
identified.
vii) Dispense 100 pl/well of 2 or 4 step dilutions of the virus to be
identified and of IPNV control virus, and allow to react with
the coated antibody to IPNV for 30 min at 37°C.
viii) Rinse once with PBST followed by three washes, allowing the
strips to soak for 3 min between washes.
ix)
Add to the wells, horse radish peroxidase (HRP)-conjugated
polyclonal antibody to IPNV; or MAb to N protein specific for
a domain different from the one of the coating MAb and
previously conjugated with biotin.
x)
Incubate 30 min at 37°C.
xi)
Rinse and wash with PBST as in step viii above.
96
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
xii) Add
the substrate
(H2O2) and
chromogen
(tetramethylbenzidine [TMB] or other approved chromogen). Stop
the course of the test when positive controls react, and read the
results.
xiii) Alternatively: add H 0
+ chromogen to those wells
containing the peroxidase conjugated antibody and proceed as
above.
2
2.
2
DIAGNOSTIC PROCEDURES FOR CONFIRMATION
OUTBREAKS
O F D?N
IN
SUSPECTED
Confirmation of IPNV can be achieved by any of the following methods:
2.1. Conventional
identification
virus
isolation
with
subsequent
serological
As Section 1.
2.2. Virus isolation with simultaneous serological identification
a)
Sampling procedures
As B . l . l . in Chapter 1 (General Information) for the selection of
fish specimens.
As B.2.1. & 2.2. in Chapter 1 (General Information) for the
selection of materials sampled.
b)
Processing of organ samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ homogenates
B.3.3. for treatment to neutralise birnaviruses (if required).
c)
Virus identification by neutralisation test
i)
Dilute organ homogenates 1:100, 1:1,000 and 1:10,000 in cell
culture medium.
ii)
Mix with equal volume of a solution of antibody to IPNV as in
1.2.a, inoculate the cell monolayers, incubate at 15°C and
monitor the fate of cell infection as in 1.2.a.
iii)
Subcultivation: if no CPE appears after one week subcultivate
the cell culture fluids of non antibody-treated controls as in
U.c.
Infectious pancreatic necrosis
97
2.3. ELISA
a)
Microplate processing
As 1.2.b. of this chapter up to point iv (inclusive).
b)
Sampling procedures
See the following sections in Chapter 1 (General Information):
B. 1.1. for the selection of fish specimens
B . 2 . 1 . & 2 . 2 . for the selection of materials sampled.
c)
Processing of organ samples
See the following sections in Chapter 1 (General Information):
B.3.1. for transportation
B.3.2. for virus extraction and obtaining of organ
homogenates.
d)
Carrying out the ELISA
i)
Set aside an aliquot of 1/4 of each homogenate in case further
virus isolation in cell culture is required.
ii)
Treat the remaining part of homogenate with 0 . 5 % PBST
(vol./vol.) as 1.2.c point v and 2 m M of phenyl methyl
sulfonide fluoride (PMSF); mix gently.
iii)
Complete the other steps of procedure 1.2.b.
REFERENCES
1.
A G I U S C , M A N G U N W I R Y O H . , JOHNSON R . H . & SMAIL D . A . ( 1 9 8 2 ) . - A
more sensitive technique for isolating infectious pancreatic necrosis virus
from asymptomatic carrier rainbow trout, Salmo gairdneri Richardson. J.
Fish Dis., 5, 2 8 5 - 2 9 2 .
2.
A H N E W. ( 1 9 7 8 ) . - Isolation and characterisation of infectious pancreatic
necrosis virus from pike (Esox lucius). Arch. Virol., 58, 6 5 - 6 9 .
3.
A H N E W. & N E G E L E R . D . ( 1 9 8 5 ) . - Studies on transmission of infectious
pancreatic necrosis virus via eyed eggs and sexual products of salmonid
fish. In: A.E. Ellis, ed. Fish and Shellfish Pathology. Academic Press,
London,261-269.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
98
4.
B U L L O C K G.L., RUCKER R.R., A M E N D D., W O L D K. & STUCKEY H.M.
( 1 9 7 6 ) . - Infectious pancreatic necrosis: transmission with iodine-treated
and non-treated eggs of brook trout (Salvelinus fontinalis). J. Fish Res. Bd
Can., 3 3 , 1 1 9 7 - 1 1 9 8 .
5.
CASTRIC J.,
BAUDIN-LAURENCIN F.,
COUSTANS M.F.
&
A U F F R E T M.
( 1 9 8 7 ) . - Isolation of infectious pancreatic necrosis virus, Ab serotype,
from an epizootic in farmed turbot, Scophthalmus maximus.
Aquaculture,
67, 1 1 7 - 1 2 6 .
6.
C H O U H.Y., L o C F . , T U N G M . C , W A N G C H . , FUKUDA H. & S A N O T.
( 1 9 9 3 ) . - the general characteristics of a birnavirus isolated from cultured
loach (Misgurnus anguillicaudatus) in Taiwan. Fish Pathol., 28, 1-7.
7.
D I X O N P.F. & H I L L B . J . ( 1 9 8 3 ) . - Rapid detection of infectious pancreatic
necrosis virus (IPNV) by the enzyme-linked immunosorbent assay. J. Gen.
Virol., 64, 3 2 1 - 3 3 0 .
8.
D O B O S P. & ROBERTS T.E. ( 1 9 8 3 ) . - The molecular biology of infectious
pancreatic necrosis virus: a review. Can. J. Gen. Microbiol,
9.
29, 3 7 7 - 3 8 4 .
D O R S O N M. & T O R C H Y C. ( 1 9 8 1 ) . - T h e influence of fish age and water
temperature on mortalities of rainbow trout, Salmo gairdneri Richardson,
caused by a European strain of infectious pancreatic necrosis virus. J. Fish
Dis.,
10.
4,213-221.
DORSON
M. & T O R C H Y
C. ( 1 9 8 5 ) .
- Experimental
transmission
of
infectious pancreatic necrosis virus via the sexual products. In: A.E. Ellis,
ed. Fish and Shellfish Pathology. Academic Press, London, 2 5 1 - 2 6 0 .
11.
EVENSEN O. & RIMSTAD E. ( 1 9 9 0 ) . - Immunohistochemical
identification
of infectious pancreatic virus in paraffin-embedded tissues of Atlantic
salmon (Salmo salar). J.Vet. Diag. Invest., 2, 2 8 8 - 2 9 3 .
12.
H I L L B . J . ( 1 9 8 2 ) . - Infectious pancreatic necrosis and its virulence. In: R.J.
Roberts, ed. Microbial Diseases of Fish (Special Publication of the Society
for General Microbiology). Academic Press, London, 9 1 - 1 1 4 .
13.
HILL B . J . ( 1 9 9 2 ) . - Impact of viral diseases of salmonid fish in the
European Community. In: T. Kimura, ed. Salmonid Diseases. Hokkaido
University Press, Sapporo, Japan, 4 8 - 5 9 .
14.
H I L L B . J . , WILLIAMS R.F. & FINLAY J. ( 1 9 8 1 ) . - Preparation of antisera
against fish virus disease agents. Dev. Biol. Stand., 49, 2 0 9 - 2 1 8 .
15.
M C K N I G H T I.J. & ROBERTS R.J. ( 1 9 7 6 ) .
- The pathology of infectious
pancreatic necrosis. I. The sequential histopathology of the naturally
occurring condition. Br. Vet. J., 132, 7 6 - 8 5 .
Infectious pancreatic necrosis
16.
99
M E L B Y H . P . , CASWELL R E N O P. & F A L K K . ( 1 9 9 4 ) . - Antigenic analysis of
Norwegian aquatic birnavirus isolates using monoclonal antibodies - with
special reference to fish species, age and health status. J. Fish Dis., 17, 8 5 91.
17.
M O R T E N S E N S.H., E V E N S E N O . , R O D S E T H O . M . & HJELTNES B . K . ( 1 9 9 3 ) . -
The relevance of infectious pancreatic necrosis virus (IPNV) in farmed
Norwegian turbot (Scophthalmus maximus). Aquaculture, 115, 2 4 3 - 2 5 2 .
18.
NAKAJIMA K . , M A E N O Y., A R I M O T O M , INOUYE K . & SORIMACHI M .
( 1 9 9 3 ) . - Viral deformity of yellowtail fingerlings. Fish Pathol., 28, 1 2 5 129.
19.
N O V O A A . , F I G U E R A S A . , PUENTES C F . , L E D O A. & T O R A N Z O A.E. ( 1 9 9 3 ) .
- Characterization of a birnavirus isolated from diseased turbot cultured in
Spain. Dis. Aquat. Organisms, 15, 1 6 3 - 1 6 9 .
20.
O K A M O T O N , SANO T., HEDRICK R.P. & F R Y E R J.L. ( 1 9 8 3 ) . - Antigenic
relationships of infectious pancreatic necrosis virus (IPNV) and European
eel virus. J. Fish Dis., 6, 1 9 - 2 5 .
21.
O K A M O T O N., TANIGUCHI N., S E N O Y. & SANO T. ( 1 9 8 4 ) . - The relation
between the change of quantities of infectious pancreatic necrosis virus in
infected rainbow trout fry and the disease process. Fish Pathol., 19, 1-4.
22.
SMAIL D.A., B R U N O D . W . , D E A R G . , M C F A R L A N E L A . & R o s s K . ( 1 9 8 2 ) .
- Infectious pancreatic necrosis (IPN) virus Sp serotype in farmed Atlantic
salmon. Salmo salar L., post-smolts associated with mortality and clinical
disease. J. Fish Dis., 15, 7 7 ' - 8 3 .
23.
24.
W O L F K . ( 1 9 8 8 ) . - Fish Viruses and Fish Viral Diseases.
University Press, Ithaca, NY, USA, 4 7 6 pp.
Cornell
W O L F K . , SNIESZKO S.F., D U N B A R C E . & P Y L E E. ( 1 9 6 0 ) . - Virus nature
of infectious pancreatic necrosis virus in trout. Proc. Soc. Exp. Biol., 104,
105-108.
Infectious salmon anaemia
101
CHAPTER
10
INFECTIOUS SALMON ANAEMIA
(No O I E number)
SUMMARY
Infectious salmon anaemia (ISA) is a viral infection of Atlantic
salmon (Salmo salar). / / is so far only reported to occur in Norway.
For more detailed information on the condition, see reference 12.
Atlantic salmon is the only susceptible fish species known to develop
the disease, but experimentally, sea trout and rainbow trout have
been shown to act as asymptomatic carriers of the disease agent.
Clinically, the initial infection appears as a systemic and lethal
condition which is characterised by anaemia, ascites,
congestions
and enlargement of the liver (dark in colour) and spleen, as well as
peritoneal petechiae. Haemorrhages in the eyes may also be seen.
Hepatocellular
degeneration
and necrosis
is a
consistent
histopathological finding. The infection is only seen in fish held in
sea water or in fish exposed to sea water. A virus, probably the ISA
agent, has recently
been isolated
and is currently
under
investigation.
The reservoirs of ISA are not known, but spread of the disease has
occurred as a result of the purchasing of subclinically
infected
Atlantic
salmon smolts; from farm to farm; and from
fish
slaughterhouses
or industries from
which organic
material
(especially blood and processing water) from ISA infected fish has
been discharged directly into sea water without further
treatment.
Few environmental factors have been identified that can be directly
linked to outbreaks of the disease. In a latent carrier
population,
various stress factors such as treatment against salmon
lice,
cestodes or infectious diseases may be followed by disease outbreaks
some 2-3 weeks later.
Due to lack of sufficient information on the ISA virus, the diagnostic
procedures for ISA are still based upon clinical,
pathological,
histopathological and haematological
changes.
The incidence of ISA has been greatly reduced by implementation of
general
legislatory
measures
regarding
movement
of
fish,
mandatory
health control, and slaughterhouse
and
transport
regulations, as well as specific measures including restrictions on
affected,
suspected
and neighbouring
farms,
epizootiological
studies, enforced sanitary slaughtering, generation segregation
fall
in/all out'), and disinfection of offals and wastewater, etc. from fish
slaughterhouses.
102
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
INTRODUCTION
Infectious salmon anaemia (ISA) is an infectious disease in Atlantic salmon
(Salmo salar).
The infective agent is possibly a virus but its real nature has not yet been
identified. ISA has so far only been described in Norway. Atlantic salmon is the
only fish species known to be susceptible to ISA, but the ISA agent may survive
for some time in sea trout (Salmo truttae) and rainbow trout
(Onchorhynchus
mykiss). The ISA agent has not been shown to survive in turbot (Psetta
maxima), bailan wrasse (Labrus berggylta), sea bass (Dicentrarchus labrax), or
cod (Gadus morhua).
DIAGNOSTIC PROCEDURES
There are no established methods for identifying the agent, and diagnosis has to
rely upon identification of typical pathological changes. None of the described
lesions are pathognomonic to ISA. The following requirements must be fulfilled
when making the diagnosis:
typical macroscopic findings and
typical histological findings and
typical haematological findings.
1.
T Y P I C A L M A C R O S C O P I C FINDINGS
Necessary finding
Dark livers
Not necessarily present in all individuals, but there
have to be some fish with dark livers. Livers can
alternatively become yellow with haemorrhagic
spots or be pale.
Supportive findings
Pale gills and heart
Result of anaemia
Ascites
Always present, early sign
Enlarged spleen
Always present, early sign
Visceral fat petecchiae
Always present
Dark foregut
Sometimes present
All these findings are typical of ISA, with some variation with regard to
severity. The more of them that are observed, the more confident one can be
regarding the diagnosis. Dark livers is an absolute criterion, as this finding is
the one most specific to ISA. Dark livers are also seen with cardiomyopathy
syndrome (CMS), which is distinguished from ISA by typical gross and
histological heart lesions.
Infectious salmon anaemia
2.
103
T Y P I C A L H I S T O L O G I C A L FINDINGS
Necessary finding
Multifocal haemorrhagic hepatic necroses, that may become confluent to give
the changes a 'zonal' appearance, leaving areas around large veins intact (late
stage of disease development).
Supportive findings
Focal congestion and dilatation of hepatic sinusoids, sometimes with
distribution as described for the necroses (early stage). Rupture of sinusoidal
endothelium with presence of erythrocytes within the space of Disse (early
sign).
Findings described as supportive are present in early stages of disease
development at haematocrit values of 15-25. The ISA-typical liver changes are
observed at heamatocrits below 10. Comparable although not identical liver
changes (usually without rupture of endothelium) may be seen in advanced
stages of CMS.
ISA is most frequently diagnosed in springtime. Mortality may be low for
months after introduction of ISA, until an 'outbreak' occurs. Necropsy and
examination of a large number of diseased or dead fish increase the probability
of detecting ISA.
Always suspect ISA when extremely dark livers are observed.
3.
T Y P I C A L H A E M A T O L O G I C A L FINDINGS
Necessary finding
Haematocrit <10.
Supportive finding
Blood smears with degenerate and vacuolised erythrocytes, and presence of
erythroblasts with irregular nuclear shape. Reduced proportion of leucocytes
relative to erythrocytes, the highest reduction among lymphocytes and
thrombocytes. Reduction of several plasma parameters, except for some
enzymes indicating liver damage, and for electrolytes (fish in seawater).
A haematocrit below 10 is not a unique finding for ISA. Fish with ulcerations
and fish suffering from erythrocytic inclusion body syndrome (ELBS), may
regularly demonstrate haematocrit values this low.
4.
VIROLOGICAL EXAMINATION
These have not yet been established for routine diagnostic purposes.
104
5.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
ZOOSANITARY PRECAUTIONS
Sanitary slaughtering
Cleaning and disinfection of farm premises and equipment
Restricted movement of live fish
Disease surveillance
Disinfection of offal/waste water from fish slaughterhouses.
REFERENCES
1.
DANNEVIG B . H . & FALK K . (1994). - Atlantic salmon, Salmo salar L.,
develop infectious salmon anaemia (ISA) after inoculation with in vitro
infected leucocytes. J. Fish Dis., 17, 183-187.
2.
DANNEVIG B . H . , FALK K . & KROGSRUD J. (1993). - Leucocytes from
Atlantic salmon, Salmo salar L., experimentally infected with infectious
salmon anaemia (ISA) exhibit an impaired response to mitogens. J. Fish
Dis., 16, 351-359.
3.
DANNEVIG B . H . , FALK K . & SKJERVE E . (1994). - Infectivity of internal
tissues of Atlantic salmon, Salmo salar L., experimentally infected with
the aetiological agent of infectious anaemia (ISA). J. Fish Dis., 17, 613622.
4.
EVENSEN 0 , T H O R U D K . E & OLSEN Y.A. (1991). - A morphological study
of the gross and light microscopic lesions of infectious anaemia in Atlantic
salmon (Salmo salar). Res. Vet. Sci., 51, 215-222.
5.
FALK K . & DANNEVIG B . H . (1995). - Demonstration of a protective
immune response in infectious salmon anaemia (ISA)-infected Atlantic
salmon (Salmo salar L). Dis. Aquatic Organisms, (in press).
6.
HASTEIN T . , ed. (1993). - Workshop on infectious salmon anaemia. Central
Veterinary Laboratory, Oslo, Norway, 55 pp.
7.
HJELTNES B., SAMUELSEN O.B. & SVARDAL A.M. (1992). - Changes in
plasma and liver glutathion levels in Atlantic salmon, Salmo salar,
suffering from infectious salmon anaemia (ISA). Dis. Aquatic
Organisms,
17,31-33.
8.
H O V L U N D T . , N Y L U N D A., W A T A N A B E K . & E N D R E S E N C.
(1994).
-
Observations of infectious salmon anaemia virus in Atlantic salmon,
Salmo salar L. J. Fish Dis., 17, 291-296.
9.
N Y L U N D A., H O V L U N D T . , H O D N E L A N D K . , NILSEN F . & LOVTK P . (1994). -
Mechanisms for transmission of infectious salmon anaemia (ISA). Dis.
Aquatic Organisms, 19, 95-100.
Infectious salmon anaemia
10.
105
OLSEN Y.A., FALK K . & REITE O . B . ( 1 9 9 2 ) . - Cortisol and lactate levels in
Atlantic salmon Salmo salar developing infectious anaemia (ISA). Dis.
Aquatic Organisms, 14, 9 9 - 1 0 4 .
11.
SPEILBERG L . , EVENSEN 0 . & DANNEVIG B . H . ( 1 9 9 5 ) . - A sequential study
of the light and electron microscopical liver lesions of infectious anemia in
Atlantic salmon (Salmo salar L ) . Vet. Pathol., (in press).
12.
THORUD K . ( 1 9 9 1 ) . - Infectious salmon anaemia. Transmission trials,
haematological, clinical chemical and morphological investigations.
Thesis for the degree of Doctor Scientarum, Norwegian College of
Veterinary Medicine.
Epizootic ulcerative syndrome
107
CHAPTER
11
EPIZOOTIC ULCERATIVE SYNDROME
(No O I E n u m b e r )
SUMMARY
Epizootic
mid and
currently
believed
Australia,
ulcerative syndrome is an epizootic infectious disease of
farmed fresh and brackish water fish. The disease is
extending through South East to South Asia and is
to be the same disease as red spot disease in Eastern
and probably the same as mycotic granuloma in Japan.
The disease occurs in almost all freshwater fish species and to a
lesser extent in brackish water fish. EUS is more prevalent
in
snakeheads fChannidae^ and barbs iPuntius spp.J. Tilapias seem to
be completely resistant. Chinese and European carp are rarely
affected.
The primary causative agent of the disease appears to be a fungus of
the genus Aphanomyces, recently termed Aphanomyces invaderis.
However, the opportunistic
bacteria Aeromonas hydrophila and
Aeromonas sobria and a number of viruses, including two distinct
groups of the rhabdoviridae and a retrovirus, may also be involved
in the pathogenesis.
Diagnosis is based on clinical signs and histological evidence
typical aggressive invasiveness of the non-sporing hyphae
fungi, in the context of heavy losses. Isolation of the fungus
its characteristic
growth profile to be used as an aid
speciation.
of the
of the
allows
to its
Control of EUS is almost impossible in the context of fish
movements since it seems to move between watersheds via wild
species. Regulation of tropical fish importation should control its
movement to islands but on land masses, spread of the disease
appears inevitable.
During outbreaks, liming of water and improvement
of water
quality, together with removal of infectedfish, is often very effective.
INTRODUCTION
Epizootic ulcerative syndrome (EUS) is an epizootic infectious disease of great
importance in wild and farmed species of fresh and brackish water fish. The
disease extended its range from Australia, where it was previously called red
spot disease (RSD), through Papua New Guinea to South East Asia and into
South Asia, where it has reached as far south as Sri Lanka. It is now
108
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
approaching Pakistan.
The most probable primary causative agent is the fungus
Aphanomyces
invaderis, which causes severe liquefactive necrosis of muscle tissue as it
invades the body, extending down into the visceral organs. Affected fish usually
also suffer from severe bacterial septicaemia, involving a variety of
opportunistic pathogens. The most frequently found bacterial pathogens are
Aeromonas hydrophila or Aeromonas sobria. A variety of parasites have also
been found among diseased fish, but their presence is inconsistent. An
associated virus infection is also frequent, involving rhabdoviridae or
myxoviridae, but retroviruses have also been isolated from affected populations.
By itself the fungus cannot normally invade fish and some co-factor such as
severe environmental stress or a virus infection is postulated as the initiating
factor to this complex and exceedingly important disease.
Almost all freshwater fish species native to a particular area and a number of
brackish water species are susceptible to EUS. Snakeheads (Channidae) and
barbs (Puntius spp.) are particularly susceptible. Chinese and European carp are
rarely affected and tilapias seem to be completely resistant.
CLINICAL SIGNS AND P A T H O L O G Y
The initial feature is usually mass mortality, with distinct dermal lesions and
ulcers, of fish in a water body. The surviving fish will show lesions and ulcers
of varying degrees of severity. They may appear as red spots, blackish burn-like
marks, or deeper ulcers with red centres and white rims. Some fish, especially
snakeheads, survive a long time with such ulcers which may erode down to the
vertebrae or so deep as to expose the brain or abdominal viscera.
Histologically, in less severely affected fish, typical non-sporulating, centrally
invasive very delicate fungi can be observed encompassed within a
granulomatous coating of epitheloid cells, relentlessly extending deep into the
visceral organs such as the kidney and liver, after it has spanned the
musculature.
DIAGNOSIS
Diagnosis is based on clinical signs and histopathology. So far. there are no
specific diagnostic tests available.
The fungus can, with some difficulty, be isolated and cultured. Apart from its
typical vulnerability to temperatures above 30°C, it is very similar to non­
pathogenic opportunistic Aphanomyces
spp., which readily contaminate the
surface of affected fish and often interfere with isolation attempts.
Experimentally, the fungus has also been shown to be pathogenic to salmonids,
after intramuscular injection of spores, with similar pathology and mortality.
Epizootic ulcerative syndrome
109
CONTROL
The control of EUS in wild populations is an impossible task. However, in small
closed water bodies, the disease can be effectively controlled by liming of water
and improvement of water quality together with removal of infected fish.
REFERENCES
1.
BALASURIYA K . S . B . , KULATHILAKE M. & SUBASINGHE R.P. (1990).
-
Preliminary investigations into the experimental transmission of epizootic
ulcerative syndrome (EUS) in fish in Sri Lanka. In: R. Hirano & I. Hanyu,
eds. Proceedings of the Second Asian Fisheries Forum, Tokyo, Japan, 1722 April 1990. Asian Fisheries Society, Manila, Philippines, 659-662.
2.
B O N D A D - R E A N T A S O M.G.,
LUMANLAN S.C., NATIVIDAD J.M.
& PHILLIPS
M.J. (1992). - Environmental monitoring of the epizootic ulcerative
syndrome (EUS) in fish from Munoz, Nurva Ecija, Philippines. In: I.M.
Shariff, R.P. Subasinghe & J.R. Arthur, eds. Diseases in Asian
Aquaculture. Fish Health Section, Asian Fisheries Society, Manila,
Philippines, 475-490.
3.
CALLINAN R.B. (1985). - Diseases of native Australian fishes. In: J.D.
Humphrey & J.S. Langdon, eds. Australian Workshop on Diseases of Fish
and Shellfish. Australian Fish Health Reference Laboratory, Benalla,
Victoria, Australia, 102-117.
4.
COATES D. (1984). - An ulcer-disease outbreak amongst the freshwater
fish population of the Sepik River system, with notes on some freshwater
fish parasites. Report No. 84-02, Dept of Primary Industries, Fisheries
Research and Surveys Branch. Port Moresby, Papua New Guinea.
5.
F A O (1986). - Report of the Expert Consultation on Ulcerative Fish
Diseases in the Asia-Pacific Region. (TCP/RAS/4508.) Bangkok,
Thailand, 5-9 August 1986. FAO, Regional Office for Asia and the
Pacific, Bangkok.
6.
FRERICHS G.N., H I L L B.J. & W A Y K . (1989). - Ulcerative disease
rhabdovirus: cell line susceptibility and serological comparison with other
fish rhabdoviruses. J. Fish Dis., 12, 51-56.
7.
FRERICHS G.N.,
M I L L A R S.D.
& ROBERTS R.J.
(1986).
-
Ulcerative
rhabdovirus in fish in Southeast Asia. Nature, 322, 216.
8.
PHILLIPS M.J. & KEDDIE H.G. (1990). - Regional Research Programme on
Relationships between Epizootic Ulcerative Syndrome in Fish and the
Environment. A Report on the Second Technical Workshop, 13-26 August
1990. Network of Aquaculture Centres in Asia, Bangkok.
110
9.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
R O B E R T S R.J., M A C I N T O S H D.J.,
THONGUTHAI K . , BOONYARATPALIN S.,
TAYAPUTCH N., PHILLIPS M.J.
& M I L L A R S.D.
(1986).
- Field
and
Laboratory Investigation into Ulcerative Fish Diseases in the Asia-Pacific
Region. Technical Report of F A O Project TCP/RAS/4508. Bangkok,
Thailand.
10.
ROBERTS R.J.,
FRERICHS
G.N.
&
MILLAR
S.D.
(1992).
-
Epizootic
ulcerative syndrome - the current position. In: I.M. Shariff, R.P.
Subasinghe & J.R. Arthur, eds. Diseases in Asian Aquaculture. Fish
Health Section, Asian Fisheries Society, Manila, Philippines, 431-436.
11.
R O B E R T S R.J., FRERICHS G.N., T H O N G U T H A I K . & CHINABUT S. (1994). -
Epizootic ulcerative syndrome (EUS) in wild and farmed fishes.
Advances in Aquaculture, 5, 207-236.
12.
Recent
R O B E R T S R.J., WILLOUGHBY L.G. & CHINABUT S. (1993). - Mycotic
aspects of epizootic ulcerative syndrome (EUS) of Asian fishes. J. Fish
Dis., 16, 169-183.
13.
SAITANU K . S . , W O N G S A W A N G S., SUNYASOOTCHAREE B. & SAHAPHONG S.
(1986). - Snakehead fish virus isolation and pathogenicity studies. In: J.L.
Maclean, L.B. Dizon & L.V. Hisillos, eds. The First Asian Fisheries
Forum. Asian Fisheries Society, Manila, Philippines, 327-330.
14.
SUBASINGHE R.P., JAYASINGHE L.P., BALASURIYA K . S . W . , KULATHILAKE
M. (1990). - Preliminary investigations into the bacterial and fungal
pathogens associated with ulcerative fish disease syndrome in Sri Lanka.
In: R. Hirano & I. Hanyu, eds. Proceedings of the Second Asian Fisheries
Forum, Tokyo, Japan, 17-22 April 1990. Asian Fisheries Society, Manila,
Philippines, 655-657.
15.
SUBASINGHE R.P. (1993). - Effects of controlled infections of Trichodina
sp. on transmission of epizootic ulcerative syndrome (EUS) to native
snakehead, Ophicephalus striatus Bloch. J. Fish Dis., 16, 161-164.
16.
TONGUTHAI K . (1985). - A Preliminary Account of Ulcerative Fish
Diseases in the Indo-Pacific Region (a Comprehensive Study based on
Thai Experiences). National Inland Fisheries Institute, Bangkok,
Thailand, 39 pp.
17.
T O R R E S J.L., SHARIFF M.
& L A W A.T.
(1990). - Identification
and
virulence screening of Aeromonas spp. isolated from healthy and epizootic
ulcerative syndrome (EUS)-infected fish. In: R. Hirano & I. Hanyu, eds.
Proceedings of the Second Asian Fisheries Forum, Tokyo, Japan, 17-22
April 1990. Asian Fisheries Society, Manila, Philippines, 663-667.
18.
T O R R E S J.L., T A J I M A K . & SHARIFF M. (1992). - Numerical taxonomy and
virulence screening of Aeromonas
spp. isolated from healthy and epizootic
Epizootic ulcerative syndrome
111
ulcerative syndrome-positive fishes. Asian Fisheries Science. 6 , 1 1 - 2 2 .
19.
WILLOUGHBY L . G . & ROBERTS R.J. ( 1 9 9 5 ) . - Improved methodology for
isolation of the Aphanomyces
fungal pathogen of epizootic ulcerative
syndrome (EUS) in Asian fishes. In: R.J. Roberts, C. Campbell & I.H.
MacRae, eds. Proceedings of the ODA Regional Seminar on Epizootic
Ulcerative Syndrome, 2 5 - 2 7 January 1 9 9 4 . Aquatic Animal Health
Research Institute, Bangkok, Thailand.
Bacterial kidney disease
113
CHAPTER
12
BACTERIAL KIDNEY DISEASE
(B408)
SUMMARY
Bacterial kidney disease (BKD) is a chronic infection with a
protracted course and an insidious nature. Fish of the Salmonidae
family are clinically susceptible,
in particular
those of the
Oncorhynchus (Pacific salmon) genus. The diagnosis depends on
increased low mortality, clinical classical and pathological
changes
associated
with the disease
as well as
histopathological
granulomatous
changes,
and
isolation
of
Renibacterium
salmoninarum in cysteine enriched media (KDM2 - SKDM) from the
lesions or from asymptomatic (latent) carriers of the pathogen.
Identification of the agent. Isolation o / R . salmoninarum may be
achieved by streaking material from diseased fish onto KDM2 or its
selective analogue SKDM plates with subsequent incubation period
at 15-18°C. Maximum incubation time reported for visible growth is
8 and 12 weeks for KDM2 and SKDM respectively, but growth is
usually obtained after 3 weeks. Identification is made according to
morphological criteria, staining reactions, catatase and oxidase
production, enzyme profiles (APl-Zym) as well as immunological
techniques (agglutination, FA T, IFA T, ELISA).
Requirements
for biological products. Several papers in the
literature report vaccination trials carried out against BKD. But no
commercial vaccine is presently available. Although some success
could be claimed experimentally, all the trials conducted in the field
resulted in rather questionable
protection.
A. D I A G N O S T I C T E C H N I Q U E S
Bacterial kidney disease (BKD) is caused by Renibacterium salmoninarum, a
coryneform Gram positive bacterium which is the sole species belonging to the
genus Renibacterium and has been reported to occur in North America, Japan,
Western Europe and Chile. Its economic importance for salmonid husbandry,
and especially for Pacific salmons (Oncorhynchus spp.), results from this wide
distribution both in fresh and saline environments, from its chronicity which
does not allow the disease to be suspected before late clinical or debilitating
manifestations, from its vertical transmission through sexual products, and from
the inefficacy of the main therapeutical compounds used in treating fish. The
special nutritive requirements and fastidious growth of the bacterium make its
diagnosis difficult. Although health control presently appears to be the best way
to control the disease, and although different methods have been suggested for
114
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
improving the detection of the agent in infected fish populations, there is not yet
general agreement on the respective value of these methods.
The overt disease only appears in advanced cases of infection, when the fish
have completed their first year of life. Exophfhalmia and abdominal distension
resulting from the impairment of the excretory function can be associated with
superficial lesions and haemorrhagia. Internal pathology is more typical.
Systemic granulomatous lesions can be found in all the organs, but develop
especially in the kidney. Greyish necrotic abscesses tend to multiply and merge
until the diffuse granulomatosis results in enlargement and necrosis of the
whole kidney, which appears swollen and bloated with irregular greyish areas.
The condition must be distinguished from proliferative kidney disease (PKD), in
which the kidney hypertrophy is not associated with any discolouration, and
from nephrocalcinosis, in which only the urinary conducts are impaired and
exhibit a white porcelain colour. In doubtful cases, microscopic observation of
smears or prints of kidney samples, stained with Gram or metachromatic dyes
(toluid blue, thionin), allows visualisation of large numbers of small bacteria
that must not be confused with the melanin granules also commonly present in
kidney tissues. In any case, clinical diagnosis provides only a suspicion of BKD,
as other Gram positive bacteria, namely lactic bacteria, have been demonstrated
to produce similar infections in salmonids. Diagnosis ought to be confirmed by
laboratory procedures.
1.
IDENTIFICATION O F T H E AGENT
Specimens can be taken from dead and preferably moribund fish showing
lesions that give reasons for suspicion of BKD. The diagnostic procedure
should follow the sequence: direct microscopy of stained smears (Gram,
PAS, Ziehl-Neelsen) from lesions in kidney or other internal organs,
possibly direct detection of antigens in tissues samples using specific
antisera against R. salmoninarum, inoculation of material onto suitable
medium for bacterial growth and identification.
Although the fastidious growth of R. salmoninarum and its special
requirements for serum and cysteine make its culture difficult, whenever
possible culturing should be used for confirmation. Special media (KDM2,
SKDM) and procedures have been developed to improve the isolation of the
bacterium, which is usually obtained after two to three weeks.
Morphological criteria, staining reactions, catalase and oxidase production,
and enzyme profiles in API-Zym panels are useful.
1.1. Isolation and bacteriological identification
a)
Sampling
Tissue samples for diagnosis and identification of R. salmoninarum
should be taken aseptically from the lesions in kidneys or other
organs. When no lesions are present the kidney should be preferred
Bacterial kidney disease
115
for sampling, but in mature females the coelomic fluid may also
represent a convenient material.
For routine controls for detecting infected individuals
population a sufficient number of fish must be sampled.
b)
in
a
Isolation
R. salmoninarum is a fastidious growing organism which requires
prolonged incubation (2 to 12 weeks at 15°C) to produce colonies.
Cysteine and serum are requisite factors, and different media or
ingredients have been proposed to improve its growth or reduce the
development of associated microorganisms. Two of these special
media are currently used:
Kidney disease medium 2 (KDM2)
L-cysteine (chlorhydrate)
Tryptone
Yeast extract
Agar
Distilled water
0.1 g
1g
0.05 g
1.5 g
100 ml
Adjust the pH to 6.5-6.8 with NaOH, distribute in flasks or tubes
and autoclave for 20 min at 120°C. Can be stored for 1 month at
4°C.
Regenerate prior to use and add 5-10% fetal calf serum.
Selective kidney disease medium (SKDM)
L-cysteine (chlorhydrate)
Cycloheximide
Tryptone
Yeast extract
Agar
Distilled water
0.1 g
0.005 g
1g
0.05 g
1g
100 ml
Adjust to pH 6.8 with NaOH and autoclave for 15 min at 121°C.
Cool to approximately 48°C, and add 10% fetal calf serum, and the
following components, previously filter sterilised (0.22 urn):
D-cycloserine
Oxolinic acid
Polymyxin B sulphate
0.00125 g
0.00025 g
0.0025 g (final concentrations)
Dishes with isolation medium are dried at room temperature for 2448 h, inoculated with 0.1 to 0.2 ml drops of infectious material, and
incubated at 15°C in plastic bags or moist chambers when the
absorption is complete.
116
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Antibiotic supplementation of KDM2 medium may reduce the
problems with fast-growing organisms (bacteria and fungi), but also
inhibit R. salmoninarum itself. This supplementation must thus be
done carefully. Another possibility is to inspect the dishes regularly
at intervals of 2-3 days, and aseptically remove the colonies
produced by fast-growing organisms. In order to respect the
viability of the KD bacterium, Evelyn recommends the preparation
of tissue suspensions in physiological saline 9 p. 1000 enriched with
peptone 1 p. 1000.
When a stock culture of R. salmoninarum is already available, it is
possible to take advantage of a "satellitism" phenomenon described
by Evelyn for accelerating the growth of the isolates. A heavy
suspension of the laboratory feeding strain is dropped onto the
centre of the plate, and the samples to be tested are inoculated in
the periphery. The growth rate and the colony size of the isolates
are noticeably increased. The growth enhancement may also be
achieved by adding 1.5% v/v sterile spent KDM2 broth to the
medium.
c)
Characteristics
After sufficient time of incubation on KDM2 and SKDM,
R. salmoninarum produces white or creamy shiny smooth, round,
raised, entire, pin point to 2 m m colonies. Bacteria from diseased
fish will on the average produce visible colonies after 2 to 3 weeks,
however up to 8 weeks have been reported for initial growth on
KDM2, and 12 weeks on the selective medium SKDM. Old cultures
may achieve a granular or crystalline appearence. Transverse
sections through such colonies will reveal the presence of Gram
positive rods in a crystalline matrix. The crystalline material is
thought to be cysteine precipitated from the medium. No growth
occurs on blood agar medium without cysteine supplement or
trypticase yeast agar. For some strains a uniformly turbid growth
occurs in broth, but for others a sediment may develop.
R. salmoninarum
appears as small (0.3-1.5 x 0.1-1 um) Gram
positive, PAS positive, asporogenous, non-motile, non-acid fast
rods, frequently in pairs, short chains or pleomorphic forms as
"Chinese letters", especially in fish tissue.
R. salmoninarum
is catalase positive and oxidase negative. Its
phenotypic characteristics have been established using API-Zym
systems and conventional tests (Tables 1 and 2). However the slow
growth of the organism does not render such tests very useful in
practice, and serological methods are more usually employed to
confirm the identity of the isolated strains.
Bacterial kidney disease
117
Table 1
Distinguishing profiles of Gram-positive bacteria morphologically similar to
R. salmoninarum, obtained with API-Zym
a-mannosidae
a-fucosidase
1
3
4
5
6
7
8
10
11
12
13
14
15
16
17
18
19
20
Actinomyces viscosus (ATCC
15987)
- - -
+
-
+
+
- - -
+
+
+
+
- -
+
- -
+
+
-
(+)
-
+
- - -
Trypsinase
Taxon (and source of strains)
Control
N-acetyl-fl-
U-glucuronidase
a-galactosidase
Phosphoamidase
Acid phosphatase
Chemotrypsinase
Cystine arylamidase
Leucine arylamidase
Valine arylamidase
Lipase (myristate)
Esterase {caprylate)
Esterase (butyrate)
Alkaline phosphatase
Character
(0
%
CO
re
o
CO
l/l
m
XI
"w
o
CJ
tA
re
8
O
O)
S
S
A P I - r / m test N o .
2
Corynebacterium a c n e s
(NCTC 7 3 7 )
9
(+)
+
C pyogenes ( N C T C 5 2 2 4 )
-
+
+
+
-
+
- -
(+)
+
-
+
(+)
Mycobacterium aqua
(Kôrmendy)
-
+
+
+
+
+
- - - -
Myc. fortuitum (Kôrmendy)
-
+
+
+
-
+
+
Myc. marinum (Kôrmendy)
-
+
+
+
-
Mycobacterium sp.
(Ashburner, S C 7 4 4 )
-
+
+
+
-
Nocardia asteroides ( A T C C
14759)
-
+
+
+
- -
+
Renibacterium salmoninarum
(48 isolates)
-
+
+
-
+
- -
Rothia dentocariosa ( A T C C
17931)
- -
+
-
+
+
-
Presumptive coryneform
(laboratory isolate 1 9 8 8 )
- -
+
-
+
- - - -
C. xerosis ( N C T C 7 9 2 9 )
Lactobacillus sp.
(pseudokidney disease, 3
+
- - -
+
isolates)
+
Modified (extracted) from Table 15.5 of reference 1 .
+
+
-
+
+
+
- - - -
+
+
+
- - - -
+
+
- - -
- -
+
+
- - - - - - - -
+
-
+
+
- - -
+
- -
+
-
+
- - -
+
- - - -
+
- - -
+
- - - -
-
+
+
+
- - - - - - - -
+
- - - -
(+)
-
118
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Table 2
C h a r a c t e r i s t i c s of Renibacterium
Character
Production of:
Acid phosphatase
Alkaline phosphatase
Butyrate esterase
Caprylate esterase
Catalase
Chymothrypsinase
Cystine arylamidase
(X—fucosidase
oc-galactosidase
P-galactosidase
P-glucosaminidase
a-glucosidase
P-glucosidase
p-glucoronidase
Leucine arylamidase
a-mannosidase
Myristate esterase
Oxidase
Trypsinase
Valine arylamidase
Response
+
+
_
+
+
_
+
+
+
-
Nitrate reduction
+
-
Degradation of:
Adenine
Aesculin
Arbutin
Casein
Chitin
+
-
Chondroitin
DNA
Elastin
Gelatin
Guanine
Hyaluronic acid
Hypoxanthine
Lecithin
_
-
salmoninarum
Character
RNA
Starch
Testosterone
Tributyrin
Tween 40
Tween 60
Tween 80
Tyrosine
Xanthine
Acid production from sugars
Response
_
_
+
+
+
-
Growth on/at:
pH 7.8
Bile salts (0.025% w/v)
Crystal violet (0.0001% w/v)
Methylene blue (0.001% w/v)
Nile blue (0.00001% w/v)
Phenol (0.025% w/v)
Potassium thiocyanate ( 1 % w/v)
Sodium chloride (1% w/v)
Sodium selenite (0.01% w/v)
Thallous acetate (0.001% w/v)
Utilisation of:
4-umbelliferyl (4MU)-acetate
4MU-butyrate
+
+
+
_
_
+ (poor)
_
»
4MU-p-D-cellobiopyranoside monohydrate
4MU-elaidate
4MU-cc-L-arabinopyranoside
4MU-2-acetamido-2-deoxy-P-Dgalactopyranoside
4MU-p-L-fucopyranoside
4PU-heptanoate
4PU-laurate
4PU-nonanoate
4MU-oleate
4MU-palmitate
4MU-propionate
+
+
_
+
+
+
+
_
+
Bacterial kidney disease
119
1.2. Antigen detection and identification by serological methods
Although some risk of cross-reactions has been reported to occur with other
bacteria the antigenic homogeneity of R. salmoninarum is a characteristic
which favours the use of specific antisera in identification procedures. Such
techniques have been applied to direct detection of the KD bacterium in
infected fish and are favoured by the existence of a soluble, heat-stable,
major antigen (antigen F) abundantly released in infected tissues. The amino
acid and DNA sequences of the main soluble antigen have been determined.
Rapid immunological tests have found broad application in monitoring
disease outbreaks and screening of brood fish to sort out the least infected
fish for propagation purposes. Slide agglutination and coagglutination are
now supplemented by direct or indirect fluorescent antibody tests (FAT and
IF AT) and by enzyme-linked immunosorbent assays (ELISA), using either
specific sera or monoclonal antibodies. A diagnostic ELISA kit approved by
the United States Department of Agriculture has recently been developed
and commercialised.
a)
Serum
It is of course necessary to obtain good antisera. When polyclonal
antisera are required this can be achieved by immunising rabbits.
One of the most effective methods to obtain sera with fair antibody
titers is the multispot intradermal schedule described in Chapter 1
(C.3.2.).
Monoclonal antibodies have also been produced against a major
surface protein which is a part of the antigen, antigen F, mentioned
above. However, the epitopes the Mabs react with are not heat
stable. Mabs are currently used in a diagnostic ELISA kit recently
commercialised (1.2.d.).
In certain cases, purified immunoglobulins, or at least enriched
fractions, may prove useful and should be prepared by classical
methods.
b) Agglutination
test
Isolated organisms can be identified quickly using slide
agglutination tests. Some colonies are gently mixed into a drop of
sterile saline, on a clean glass slide, and a drop of antiserum is
added. The agglutination of the bacteria can be observed by
comparing a similar suspension in normal rabbit serum as control.
The appropriate dilution of reacting serum will have been
previously determined by testing a control R. salmoninarum strain
in twofold dilutions of this serum.
In order to improve and facilitate the agglutination, coagglutination
120
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
using Staphylococcus
aureus (Cowan I strain) sensitised with
specific immunoglobulins has been described.
Staphylococcus,
cultivated 24 h at 37°C in liquid medium, is washed and
successively treated by formalin 0.5% and heating 30 min at 100°C.
The suspension is adjusted to 10% (v/v) in phosphate buffer and
added to 0.1 volume of mti-Renibacterium
Ig. Incubation is 3 h at
25°C, with periodic stirring. The sensitised bacteria are then
centrifuged and adjusted to the initial volume after washing
3 times. Agglutination tests are performed on glass slides
maintained in moist chambers and read after 30, 60 and 120 min.
According to the authors this test provides better and quicker
results when the antigen to be tested is heated 30 min at 100°C. It
appears, therefore, that the specific reaction is supported mainly by
the heat stable antigen known as antigen F. As antigen F is soluble
and widely released in infected tissues coagglutination has been
applied to direct detection of R. salmoninarum
in kidney tissue
preparations. Despite its low requirements in materials it does not
seem to ensure as good a sensitivity as immunofluorescence or
ELISA tests.
c)
Immunofluorescence
Direct and indirect immunofluorescent antibody tests (FAT, IF AT)
have been commonly used in demonstrating the presence of the
bacterium in tissues. In the first case, rabbit or goat anti-BK
bacterium immunoglobulins conjugated to fluorescein are used. In
the second case, heterospecific Igs prepared against the anti-BKD
serum are labelled. Tissue smears prepared from infected kidney are
air-dried and heated 2 min at 60°C before being incubated with the
reagents and observed microscopically through UV light. This
technique has proved sensitive enough to permit detection of carrier
fish, as well as to control eggs and ovarian fluids of mature fish
selected as breeders. However, an interlaboratory comparison of the
IF AT revealed that reproducibility was poor for the lowest levels of
infection and moreover several infections were missed as shown by
parallel culture. To obviate risks of cross-reaction with other
coryneform bacteria the alternative use of monoclonal antibodies
against specific determinants has been recommended.
d) Enzyme-linked
immunosorbent
assay
(ELISA)
ELISA can be used for the detection of soluble antigen extracted
from infected tissue and is probably the most sensitive method. It
allows testing of large numbers of samples and is specially well
adapted to the requirements of health certification. The most
common technique is the double antibody sandwich one: specific
antibodies are coated onto the polystyrene or P V C surface of
Bacterial kidney disease
121
microplate wells, and samples containing the antigen, conjugate
serum, and substrate are successively added.
Two different ELISA procedures for soluble antigens have been
used. One uses polyclonal antibodies reacting with heat stable
antigenic determinants, while the other uses monoclonal antibodies
towards heat labile antigenic determinants. The heat treatment
limits cross reactions, but not completely, as shown by cross
reactions with feather meal used in fish feed. The monoclonal
antibodies appear very specific, but the samples must be kept cold
or frozen after sampling. Diseased and subclinically infected fish
give ELISA reactions that are clearly distinct from uninfected fish.
However, the exact positive threshold is difficult to set as
representative non-infected fish, for sdetermining
normal
background variation, are not always easy to find.
Problems linked to cross-reactions are the same as in
immunofluorescence and could be solved definitively using
monoclonal antibodies. A diagnostic ELISA kit approved by the US
Department of Agriculture has been developed by DiagXotics Inc.
(126 Old Ridgefield Rd, Wilton, CT 06897, USA). All the materials
and reagents are supplied to perform quantitative tests which run
exactly as previously described, except for two modifications:
microplates are sold ready for use, coated with a first monoclonal
antibody. After sample distribution a second Mab of different
specificity and conjugated to biotin is added. This allows the
sensitivity to be improved by using the avidin-biotin system,
streptavidin being conjugated to the horse radish peroxidase added
in the subsequent step. Control antigen solutions are also supplied
to allow a standard curve to be established.
2. Serological tests
Serological tests are not used routinely for the diagnosis of BKD. In fact, the
development of the antibody response does not correlate clearly with the
course of the infection, and several authors disagree with the adoption of
serology as a health inspection method.
B. R E Q U I R E M E N T S F O R B I O L O G I C A L P R O D U C T S
No commercial vaccines against BKD are available, but they are greatly needed
as R. salmoninarum responds poorly to medical treatment. There is evidence
that Renibacterium under some conditions elicits an immune response in fish,
and there are some reports of vaccination in the literature. The protective ability
of a vaccine is however questionable and basically one of the problems is the
intracellullar nature and vertical transmission of the agent. This will require a
much better understanding of the pathogenesis of R. salmoninarum, and how it
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
122
interacts with the immune system of the fish. It may only be possible to
vaccinate non-exposed fish, as exposed and subclinically infected fish may not
respond to vaccination. When developed, suitable vaccines should give
protection to vaccinated fish throughout the whole lifecycle in fish farms under
normal conditions.
For the moment, health control based on specific detection strategies and
elimination of carrier or diseased fish still represents the best way to combat
BKD.
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E M B L E Y T.M., GOODFELLOW M., M I N N I K I N D.E. & AUSTIN B. (1983). -
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EVELYN T.P.T. (1971). - The agglutinin response in sockeye salmon
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EVELYN T.P.T. (1977). - A n improved growth medium for the kidney
disease bacterium and some notes on using the medium. Bull. Off. Int.
Epiz., 87, 511-513.
13.
EVELYN T.P.T. (1978). - Sensitivities of bacterial kidney disease detection
methods with special remarks on the culture method. In: Proceedings of
the Joint 3rd Biennal Fish Health Section, American Fisheries Society, and
9th Midwest Fish Disease Workshops, Kansas City, 1-2.
14.
EVELYN T.P.T, B E L L G . R . , P R O S P E R I - P O R T A L. & K E T C H E S O N J.E. (1989).
- A simple technique for accelerating the growth of the kidney disease
bacterium Renibacterium
salmoninarum
on a commonly used culture
medium (KDM2). Dis. Aquat. Org., 7, 231-234.
15.
FIEDLER F. & DRAXL R . (1986). - Biochemical and immunochemical
properties of the cell surface of Renibacterium
salmoninarum.
J.
Bacteriol., 168, 799-804.
16.
FRYER J.L. & SANDERS J.E. (1981). - Bacterial kidney disease of salmonid
fish. Ann. Rev. Microbiol., 35, 273-298.
17.
GETCHELL R . G . , ROHOVEC J.S. & F R Y E R J.L. (1985). - Comparison of
Renibacterium
20, 149-159.
18.
salmoninarum
isolates by antigenic analysis. Fish
GOODFELLOW M., EMBLEY T.M. & AUSTIN B. (1985). -
taxonomy and emended description of Renibacterium
Gen. Microbiol., 131, 2739-2752.
19.
Pathol.,
Numerical
salmoninarum.
GUDMUNDSDOTTIR S., BENEDIKTSDOTTIR E. & HELGASON S. (1993).
J.
-
Detection of Renibacterium salmoninarum in salmonid kidney samples: a
comparison of results using double sandwich ELISA and isolation on
selective medium. J. Fish Dis., 16,185-195.
124
20.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
H s u H-M., B O W S E R P.R. & SCHACHTE J R J.H. (1991). - Development and
evaluation of a monoclonal-antibody-based enzyme-linked immunosorbent
assay for the diagnosis of Renibacterium
salmoninarum
infection. J.
Aquat. Anim. Health, 3, 168-175.
21.
KIMURA T. & YOSHIMIZU M. (1981). - A coagglutination test with
antibody-sensitized staphylococci for rapid and simple diagnosis of
bacterial kidney disease (BKD). In: Fish biologies: sérodiagnostics and
vaccines. Dev. Biol. Stand, S. Karger, Basel, 49, 135-148.
22.
L A I D L E R L . A . (1980). - Detection and identification of the bacterial kidney
disease (BKD) organism by the indirect fluorescent antibody technique. J.
Fish Dis., 3, 67-69.
23.
L E E E.G. & GORDON M.R. (1987). - Immunofluorescence screening of
Renibacterium salmoninarum in the tissues and eggs of farmed chinook
salmon spawners. Aquaculture, 65, 7-14.
24.
LEEK S. (1988). - Sonicating kidney tissue to enhance liberation of
Renibacterium salmoninarum cells. Am. Fish Health Sect. Newslet., 16, 4.
25.
M C C A R T H Y D.H., C R O Y T.R. & A M E N D D.F. (1984). - Immunization of
rainbow trout, Salmo gairdneri Richardson, against bacterial kidney
disease: preliminary efficacy vaccination. J. Fish Dis., 7, 65-71.
26.
P A S C H O R J . , E L L I O T D . G , F O W L E R L . G . & M C K I B B E N C L . (1991). - Use
of feather meal as an alternate protein source in salmon feed and its effect
on mortality of chinook salmon due to infection by
Renibacterium
salmoninarum. In: Proceedings 14th Annual American Fisheries Society,
Fish Health Section Meeting. Newport, Oregon, p. 2 1 .
27.
PASCHO R.J. & MULCAHY D. (1987). - Enzyme-linked immunosorbent
assay for soluble antigen of Renibacterium salmoninarum, the causative
agent of salmonid bacterial kidney disease. Can. J. Aquat. Sci., 44, 183191.
28.
PATERSON W . D . , DESAUTELS D. & W E B E R J . (1980). - The
immune
response of Atlantic salmon, Salmo salar L . , to the causative agent of
bacterial kidney disease Renibacterium salmoninarum. J. Fish Dis., 4, 99111.
29.
SANDERS J.E. & FRYER J . L . (1980). - Renibacterium salmoninarum gen.
nov., the causative agent of bacterial kidney disease in salmonid fishes.
Int. J. Syst. Bacteriol., 30, 496-502.
30.
SANDERS J . E . & BARROS M . J . (1986). - Evidence by the fluorescent
antibody test for the occurrence of Renibacterium salmoninarum
among
salmonid fish in Chile. J. Wildl. Dis., 22, 255-257.
Bacterial kidney disease
31.
125
TURAGA S.D., WIENS G . D . &
Renibacterium
salmoninarum
KAATTARI
S.L.
(1987).
-
Analysis
of
antigen production in situ. Fish Pathol., 22,
209-214.
32.
W I E N S G . D . & KAATTARI S . L . ( 1 9 8 9 ) . - Monoclonal antibody analysis of
common surface protein(s) of Renibacterium
24, 1-7.
salmoninarum.
Fish
Pathol.,
Edwardsiellosis
127
CHAPTER
13
ENTERIC SEPTICAEMIA OF CATFISH
(Edwardsiellosis)
(B414)
SUMMARY
Enteric septicaemia
of catfish or edwardsiellosis
is a bacterial
disease first described in channel catfish in the USA and caused by
Edwardsiella ictaluri, a Gram negative motile bacillus belonging to
the family Enterobacteriaceae.
There are indications that E. ictaluri
consists of more than one serotype, recent isolates from
Thailand
being different from most of the US isolates.
All strains of E. ictaluri are reported virulent and cause high
mortality in warm water fish species, in particular fishes
belonging
to the family Ictaluridae. This disease condition is also known as
enteric
septicaemia.
Identification of the agent: The identification ofE. ictaluri is based
on the isolation of the causative agent and characterisation
by
biochemical tests. E. ictaluri can easily be differentiated from E.
tarda by its inability to produce indole and H2S. E. tarda produces
both. Additionally the two species do not cross react serologically.
Serological tests.
For rapid results, and due to its antigenic
homogeneity,
E. ictaluri may be identified by means of slide
agglutination, the indirect fluorescent antibody technique (IFA T), or
the enzyme-linked immunosorbent assay (ELISA). Serologically, the
antibody response against the agent may be assessed by direct
quantitative
microagglutination,
passive
haem-agglutination
or
ELISA using polyclonal or monoclonal reagents. However the ELISA
and certain other techniques are still limited to specialised
research
laboratories.
Requirements
for biological
products:
Since E. ictaluri is
immunogenically
active, production
of vaccines soon
appeared
feasible, and indeed, a commercial bacterin has appeared on the US
market. The vaccine consists of inactivated bacteria and may be
administered
by immersing
young fish
in properly
diluted
preparations. Boosters can be given orally prior to the disease peak
of autumn. However, international standards are not yet available.
128
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
A. D I A G N O S T I C T E C H N I Q U E S
Enteric septicaemia of catfish (ESC) is caused by the bacterium Edwardsiella
ictaluri which belongs to the Enterobacteriaceae family. Since its first
recognition in 1976 in channel catfish (Ictalurus punctatus) from Alabama and
Georgia, USA, ESC has become the foremost bacterial disease problem in the
commercial catfish industry in the South Eastern United States. However, E.
ictaluri has also been reported from Ciarías batrachus in Thailand and from
several ornamental species, and the susceptibility of other species including
salmonids has been shown experimentally. There are indications that E. ictaluri
consists of more than one serotype, recent isolates from Thailand being different
from most of the US isolates.
The natural reservoir of E. ictaluri seems to be the intestine of the fish, from
which the faeces disseminate the organism into the environment. E. ictaluri can
survive in pond bottoms for an extended period of time and these may thus be a
source of infection. The disease occurs only within a limited temperature range,
from 18 to 28°C. This results in seasonal fluctuations, spring and autumn being
the critical periods. Other environmental factors (water quality, organic
compounds, stocking density and stress factors) are known to modulate the
virulence of the agent. In spite of these characteristics E. ictaluri is generally
considered to be a true obligate parasite.
The clinical signs may differ according to the fish species. Two clinical forms of
ESC have been described. Most frequently the infection seems to begin in the
olfactory sacs through the nasal route, and to progress slowly upwards to
generate granulomatous inflammation in brain tissues. This chronic meningo­
encephalitis can account for behavioural manifestations, with alternating
listlessness and chaotic swimming. In typical cases, "hole in the head", a deep
erosion of the dorso-cranial part of the head which may extend to the frontal
bone is observed. Although general infection can result from such chronic
forms, it is more frequently associated with acute septicemia following enteritic
infection. Then, as in many other bacterioses, skin petechia and haemorrhages
are observed around the mouth, on the throat and at the base of the fins.
Anaemia and exophthalmia are frequent. Internally, haemorrhages and necrotic
foci are scattered in the liver and other internal organs. Enteritis, systemic
oedema, accumulation of ascitic fluid in the general cavity and enlargement of
the spleen are not really specific. Histological examination reveals a systemic
infection of all organs and skeletal muscles, with diffuse granulomatosis.
The epidemiology and pathogenesis of the disease are poorly documented.
However, there are indications that E. ictaluri is able to survive in sediments for
some months. The natural reservoir of the organism seems to be the intestine of
fish, from which it disseminates with faeces into the environment.
Disease outbreaks have been controlled by management procedures to reduce
stress, or by feeding the catfish with oxytetracycline (Terramycin® ) at the rate
Edwardsiellosis
129
of 25-30 mg/kg fish/day for 10 days. Potentiated sulphonamides have also been
used to control E. ictaluri outbreaks, but resistances to these products appeared
recently. It seems that vaccination might now be tried as a preventive treatment.
1.
IDENTIFICATION O F T H E AGENT
1.1. Sampling
Bacteriological sampling from freshly dead or moribund fish, taken
aseptically from either spleen, liver or kidney tissue, can be streaked
onto blood agar plates, brain heart infusion agar or nutritient agar
plates. The bacterium grows rather slowly but does not require special
nutrients. Optimal temperature for incubation is 25°C.
For routine sampling of fish populations, the rules are the same as
those defined for other fish infectious agents (cf. Chapter 1 Part B.1.2.
and ref. 13).
1.2. Characteristics
The bacterium grows rather slowly but does not require special
nutrients, and its biocheminal characteristics appear homogeneous in
all isolates.
Following incubation at 26°C for 48 hours, E. ictaluri appears as
smooth, circular (1-2 mm diameter), slightly convex with entire edges,
non-pigmented colonies. It is a Gram negative rod, measuring 0.75-2.5
pm, weakly motile by means of a peritrichous flagellation. This
motility generally fails to be observed at 37°C. Growth at a higher than
optimal temperatures may result in development of avirulent strains.
After having obtained the bacterium it should be identified by
biochemical and serological characteristics. Table 1 shows some of the
characteristics of the species and biogroups of the genus Edwardsiella
as given in Bergey's Manual of Systematic Bacteriology. The bacterium
prefers to grow in low temperatures (25-30°C), and although
biochemical characteristics can be studied at 37°C, variations can be
found, namely in the motility and the production of gas from formate
and glucose. E. ictaluri is biochemically less active than the other
Edwardsiella species, but it appears homogeneous. No clearcut biotype
variation is detected. E. ictaluri and E. tarda may be differentiated
from each other biochemically by the production of indole and
hydrogen sulphide (H2S). E. tarda produces both, while E. ictaluri
does not. The capacity of all isolates of E. ictaluri of degrading
chondroitin sulfate might be an important virulence factor in the
formation of the "hole in the head" lesions in affected fish.
130
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Table 1
Differentiation of the species and biogroups of the genus
E. tarda
Wild type
E. hoshinae
Edwardsiella*
E. ictaluri
Biogroup 1
Characteristic acid
production from:
D-Mannitol
-
+
+
-
Sucrose
-
-<-
+
-
Trehalose
-
-
-1-
-
L-Arabinose
-
+
(-)
-
Tetrathionate reduction
+
-
+
Malonate utilisation
-
-
+
-
Indole production
-t-
-
-
H2S production in TSI
+
-
Motility
-t-
-t-
Citrate (Christensen's)
+
+
-
_**
(+)
-
* Bergey's Manual of Systematic Bacteriology
** Weakly motile according to Hawkes, 1979.
1.3. Detection of the bacterial antigens by serological methods
In addition to well defined biochemical characteristics, there is a strong
homogeneity in the antigenic structure of the species, which does not
share any serological relationship with E. tarda. This confers a real
value to serological methods for achieving rapid diagnosis. Slide
agglutination with specific antisera against E. ictaluri as well as
fluorescent antibody techniques and enzyme immunoassay methods
have been used to provide confirmatory diagnosis. Specific sera are
obtained using formalin killed bacterins to immunise rabbits according
to classical standards, but in certain cases monoclonal antibodies may
also be be used.
a) Agglutination
test
Isolated colonies are gently mixed into a drop of sterile saline on a
clean glass slide, and a drop of antiserum is added. The
agglutination of the bacteria can be evaluated comparing a similar
suspension in normal rabbit serum as control. The appropriate
dilution of reacting serum will have been previously determined by
testing a control E. ictaluri strain in twofold dilutions of this serum.
131
Edwardsiellosis
b) Specific
immunofluorescence
The indirect fluorescent antibody technique (JJFAT) may be
employed on bacterial smears, or smears from infected organs, for
rapid confirmation of a clinical diagnosis. Smears are air-dried and
heated 2 min at 60°C before being flooded and incubated for 5 min
with specific rabbit antibody. They are washed in phosphate
buffered saline pH 7.2, flooded for 5 min with heterospecific
immunoglobulins prepared against the anti-ii. ictaluri rabbit serum,
and conjugated with fluorescein isothiocyanate. After rinsing, the
slides are observed microscopically through UV light. Improvement
of the technique by the use of monoclonal antibody has also been
reported.
c) Enzyme
immunoassay
An enzyme immunoassay adapted to the direct identification of the
bacteria in tissues smears from infected fish has been described.
Smears are prepared as for IF AT, and the first steps are similar. But
the second incubation step (5 min) uses heterospecific immraioglobulin against rabbit antiserum, conjugated to horseradish
peroxidase. A third incubation step with a substrate (DMOB Sigma)
is performed for 10 min, and after washing and drying the smears
are mounted in buffered glycerin and microscopically observed. It
may happen that smears are too thick and produce non-specific
retention of the staining. This can be solved by rinsing the smear
again for 1 or 2 min in IN HC1.
2.
SEROLOGICAL TESTS
Although antibody detection tests are rarely used for routine dïajpiœttic
purposes and are not yet approved as official procedures, they could be off
great value for the mass control of large numbers of fish required wifli the
development of health control policies. This is supported by the specificity off
the bacterium and the demonstration of circulating antibodies against E.
ictaluri in the serum of fish recovering from or surviving the disease.
2.1. Microagglutination test
Direct microagglutination, performed in 96-well reund-bcr.on»
microplates as described for other bacterial pathogens, can provide
quick quantitative data at minimal cost when high sensitivity is not
specially required. It just needs a formalin-killed bacterial suspension
prepared according to the usual techniques (i.e. formalin 3.5 p . 1000
overnight) and adjusted at 5 x 1 0 ufc/ml. Twofold dilutions of the sera
are made in physiological saline, so that the final volume is
0.25 ul/well. Antigen is added at 75 ul and the plates are incubated 2 h
at 37°C and overnight at 4°C before reading. Controls include a rabbit
8
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
132
reference serum of previously established titre and antigen incubated in
saline.
2 . 2 . Passive haemagglutination
This technique has been described using E. ictaluri LPS ( 1 mg/ml in
PBS, pH 7 . 2 ) passively coated on human O red blood cells at 4 % .
Tested sera must be absorbed with group O human red blood cells to
remove non-specific agglutinins, and heated 3 0 min at 4 5 ° C before
dilutions are done in veronal buffered saline. Coated blood cells are
used at 1%. Incubation is 6 h at room temperature and overnight at
4°C. Controls include coated and uncoated red cells in buffer and
coated cells in serum.
2 . 3 . Indirect enzyme-linked immunosorbent assay (ELISA)
The ELISA method has also been proposed, but should be standardised
to be adopted on a large scale. The best method seems to be to use
soluble major antigen obtained by sonication to coat the microplate
wells. The subsequent steps consist of diluting sera to be tested, adding
conjugated anti-fish immunoglobulin serum (monoclonal antibody can
be used), and the substrate. This technique could prove useful to follow
the antibody response to E. ictaluri in fish subjected to vaccination
programmes. But it needs to be further evaluated for accuracy and
reproducibility of the results.
B. REQUIREMENTS FOR BIOLOGICAL PRODUCTS
E. ictaluri is known to induce an antibody response after natural disease or
active immunisation. The possibility of protecting channel catfish populations
by vaccination has been comprehensively studied, and reviewed by Plumb in
1 9 8 8 . However, no commercial vaccine against ESC was available untill 1 9 9 1 ,
when an inactivated bacterin was licensed in the USA. The recommended
procedure for use is in two steps: 1 ) immersion vaccination of young fish ( 1 litre
of bacterin for 1 0 0 kg fish); 2 ) oral booster just prior to the critical period of
autumn, feeding a medicated diet at 3 % body weight for 5 days. The vaccine
was marketed after three years of successful field trials, but it will be of course
necessary to wait for large scale results before assessing its value in an anti-ESC
strategy. Although it is generally produced and harvested according to norms
given by the US Food and Drug Administration for the use of vaccines for fish,
no international directives or standards have as yet been published.
REFERENCES
1.
A I N S W O R T H A.J.,
CAPLEY G . , WATERSTREET P. & M U N S O N D. ( 1 9 8 6 ) . -
Use of monoclonal antibodies in the indirect fluorescent antibody
technique (IFA) for the diagnosis of Edwardsiella ictaluri. J. Fish Dis., 9,
439-444.
Edwardsiellosis
2.
133
B A X A D.V., G R O F F , J . M . WISHKOVSKY A. & HEDRICK R.P. ( 1 9 9 0 ) .
Susceptibility of nonictalurid fishes to experimental
Edwardsiella ictaluri. Dis. Aquat. Org., 8, 1 1 3 .
3.
infection
-
with
BERTOLINI J . M . , CIPRIANO R., P Y L E S.W. & MCLAUGHLIN J.A. ( 1 9 9 0 ) . -
Serological investigation of the fish pathogen Edwardsiella ictaluri, cause
-of enteric septicemia of catfish. J. Wildl. Dis., 26, 2 4 6 - 2 5 2 .
4.
FARMER J . J . & M C W O R T H E R A.C. ( 1 9 8 4 ) . - Genus Edwardsiella
Ewing
and McWorther 1 9 6 5 . In: Bergey's Manual of Systematic Bacteriology.
N . R . Krieg & J.G. Holt, eds. William & Wilkins, Baltimore, Maryland, pp.
486-491.
5.
HAWKE J.P. ( 1 9 7 9 ) . - A bacterium associated with the disease of pond
cultured channel catfish, Ictalurus punctatus. J. Fish. Res. Board Can., 36,
1508-1512.
6.
HAWKE J.P., MCWHORTER A . C ,
STEIGERWALT A.G. & B R E N N E R D.J.
( 1 9 8 1 ) . - Edwardsiella ictaluri sp. nov., the causative agent of enteric
septicaemia of catfish. Int. J. Syst. Bacteriol., 31, 3 9 6 - 4 0 0 .
7.
KASORNCHANDRA J., ROGERS W.A. & PLUMB J.A. ( 1 9 8 7 ) . -
Edwardsiella
ictaluri from walking catfish, Ciarías batrachus L., in Thailand. J. Fish
Dis.,
8.
10, 137-138.
KLESIUS P.M. & H O R S T M . N . ( 1 9 9 1 ) . - Characterisation of a major outer-
membrane antigen of Edwardsiella
ictaluri. J. Aquat. Anim. Health, 3,
181-187.
9.
KLESIUS P., JOHNSON K., D U R B O R O W R. & VINITNANTHARAT S. ( 1 9 9 1 ) . -
Development and evaluation of an enzyme linked immunosorbent assay
for catfish serum antibody to Edwardsiella ictaluri. J. Aquat.
Anim.
Health, 3, 9 4 - 9 9 .
10.
MIYAZAKI T., & PLUMB J.A. ( 1 9 8 5 ) . - Histopathology of
ictaluri in channel catfish, Ictalurus punctatus
Edwardsiella
(Rafinesque). J. Fish Dis.,
8, 3 8 9 - 3 9 2 .
11.
NEWTON
J . C . , W O L F E L.G.,
GRIZZLE J . M . &
P L U M B J.A. ( 1 9 8 9 ) .
-
Pathology of experimental enteric septicaemia in channel catfish, Ictalurus
punctatus (Rafinesque), following immersion exposure to Edwardsiella
ictaluri. J. Fish Dis., 12, 3 3 5 - 3 4 7 .
12.
PLUMB J.A. ( 1 9 8 8 ) . - Vaccination against Edwardsiella ictaluri. In: Fish
vaccination. A.E. Ellis, ed. Academic Press, pp. 1 5 2 - 1 6 1 .
134
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
13.
PLUMB J.A. & KLESIUS P. (1988). - An assessment of the antigenic
homogeneity of Edwardsiella ictaluri using monoclonal antibody. J. Fish
Dis., 11, 499-510.
14.
ROGERS W . A . (1981). - Serological detection of two species of
Edwardsiella
infecting catfish. In: International symposium on fish
biologies: sérodiagnostics and vaccines. Dev. Biol. Stand., 49, 169-172.
15.
SAEED M.O. & PLUMB J. A. (1987). - Serological detection of Edwardsiella
ictaluri Hawke lipopolysaccharide antibody in serum of channel catfish,
Ictaluruspunctatus
Rafinesque. J. Fish Dis., 10, 205-209.
16.
SHOTTS E.B., BLAZER V.S. & W A L T M A N W . D . (1986). - Pathogenesis of
experimental Edwardsiella ictaluri infection in channel catfish
punctatus). Can. J. Fish. Aquat. Sci., 43. 36-42.
17.
WALTMAN W . D . , SHOTTS E.B. & H s u T . C . (1986). -
characteristics of Edwardsiella
101-104.
18.
ictaluri. Appl. Environm.
(Ictalurus
Biochemical
Microbiol.,
51,
WATERSTRAT P., AINSWORTH J. & CAPLEY G. (1989). - Use of an indirect
enzyme linked immunosorbent assay (ELISA) in the detection of channel
catfish, Ictalurus punctatus
(Rafinesque), antibodies to
Edwardsiella
ictaluri. J. Fish Dis., 12, 87-94.
135
Piscirickettsiosis
CHAPTER
14
PISCIRICKETTSIOSIS
(No OIE number)
SUMMARY
Piscirickettsiosis is a disease in salmonids caused by Piscirickettsia
salmonis and first described in farmed coho salmon (Oncorhynchus
kisutch, Walbaum). The disease was first described from Chile in
1989. P. salmonis is placed in the family Rickettsiaceae.
Identification of the agent: The identification of P. salmonis is based
on isolation on CHSE-214 cell line of the causative agent with
subsequent testing regarding characteristics for rickettsiae.
The
organism may easily be distinguished
from Chlamydia, not
demonstrating
the characteristic chlamydia developmental
cycle.
The identity may also be confirmed by means of serological tests.
Serological tests: For a rapid result, the identity of P. salmonis
isolated in cell cultures or observed in smears from
pathological
material may be confirmed by means of the fluorescent antibody test
(FAT)
or immunohistochemical
methods
with
type-specific
antiserum.
Control measures: The implementation of hygienic measures and
control policy rules are the only control methods
currently
available.
INTRODUCTION
Piscirickettsiosis is a septicaemic condition in salmonids. The causative agent of
the disease is a Rickettsia named Piscirickettsia salmonis. The disease has so
far been described from Canada, Chile, Ireland, and Norway.
P. salmonis has been detected in coho salmon (Oncorhynchus kisutch), chinook
salmon (O. tschawytscha),
Sakura salmon (O. masou), rainbow trout (O.
mykiss), pink salmon (O. gorbuscha), and Atlantic salmon (Salmo salar).
The first evidence of disease may be the appearance of small white lesions in
the skin or shallow haemorrhagic ulcers. Affected fish appear dark and
lethargic, hanging at the net sides. The major gross pathological changes are
gill pallor, peritonitis, ascites, slightly enlarged spleen, swollen grey kidneys
and liver with large pale necrotic lesions. Mortality is reported to be 30-90% in
Pacific salmon.
The means by which the disease is transmitted is not understood. It may be
transmitted horizontally in sea water, or vector-borne. The disease has primarily
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
136
been repoted as a problem in marine farming, but has also been described from
fresh water farms.
Treatment with anti-bacterials may have some benefit, but is not optimal for
controlling the disease. Eggs may be disinfected. Stamping out procedures may
be used.
DIAGNOSTIC PROCEDURES
The monitoring for and diagnosis of piscirickettsiosis is based upon two
methods: isolation of the rickettsial agent in cell culture, or detection in
acridine orange (AO) or Giemsa stained tissue smears, followed by the FAT for
further identification.
Infected fish material suitable for virological examinations is:
-
d u r i n g overt infection : kidney and liver.
Sampling procedures: See Chapter 1 Part B.
1.
STANDARD MONITORING METHODS FOR PISCIRICKETTSIOSIS
1.1. Isolation of P. salmonis in cell culture
Cell line(s) to be used: CHSE-214 without antibiotics added.
a) Preparation
of tissue
i)
The kidney must be aseptically removed and transferred to a
sterile container. No antibiotics can be used at any step in the
isolation procedure. Tissues must be kept at 4°C or on ice until
processed, and must not be frozen.
ii)
Kidney tissue should be homogenised at 1:20 in antibiotic-free
balanced salt solution (BSS) then, without centrifugation,
further diluted 1:5 and 1:50 in antibiotic-free BSS for
inoculation onto cell cultures. Final dilutions for use are 10"
and 10" .
2
3
b)
Inoculation
of cell
2
monolayers
3
i)
A 10" and 10" dilution of the organ homogenates should be
inoculated on to cultured cells. The CHSE-214 cell line should
be used for isolation. Cultures must be maintained in
antibiotic-free medium.
ii)
The diluted homogenate can be inoculated directly
(0.1
137
Piscirickettsiosis
ml/culture) into the antibiotic-free culture medium overlaying
the cells.
iii)
The cell cultures must be incubated at 15-18°C for 28 days
and observed for the appearance of cytopathic effect (CPE).
The rickettsial CPE consists of plaque-like clusters or rounded
cells. With time, the CPE progresses until the entire cell sheet
is destroyed.
iv)
If no CPE occurs (except in the positive control) cultures
should be incubated at 15-18°C for an additional 14 days.
1.2. Identification of P. salmonis in cell culture
a) Fluorescent
antibody test (FA T)
i)
The identity of rickettsiae isolated in cell culture or observed
in AO stained smears must be verified by serological methods,
e.g. FAT.
ii)
Fluid from cell cultures showing extensive CPE can be spotted
directly onto microscope slides and tested in the FAT.
iii)
Tissue smears to be examined by FAT must be freshly
prepared or stored at <-20°C. Preliminary results indicate that
storage at temperatures >-4°C is not advisable.
1.3. Identification of P. salmonis in acridine orange tissue smears
a) Preparation
of acridine orange (AO) stain (Lauer et. ai,
1981)
The following stain formulation should be used: Add 20 mg AO
powder to 190 ml of sodium acetate buffer (100 ml 1 M
C H C 0 N a . 3 H 0 and 90 ml 1 N HC1). Adjust the pH to 3.5 with 1
N HC1. Store in a brown bottle at ambient temperature.
3
2
b) Preparation
2
and observation of slides
i)
Smears or impressions of the kidney, liver and spleen should
be prepared, air dried, and fixed for five minutes in absolute
methanol. The slides should then be flooded with AO stain for
two minutes, rinsed with tap water, and air dried.
ii)
Slides should be observed under oil with a fluorescence
microscope enquipped with the appropriate filter blocks.
iii)
In AO stained preparations, the rickettsiae can appear bright
red-orange or green in colour. Individual organisms are 0.5-
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
138
1.5 Ltm in diameter. They are pleomorphic, predominantly
coccoid but also appear as rings, and pairs of curved rods are
frequently observed.
2.
D I A G N O S T I C P R O C E D U R E S F O R C O N F I R M A T I O N O F P I S C I R I C K E T T S I O S I S IN
SUSPECTED O U T B R E A K S
Confirmation of piscirickettsiosis can be achieved by any of the following
methods:
2 . 1 . Conventional isolation of P. salmonis in cell culture with subsequent
serological identification as described in Section 1.
2 . 2 . Detection in tissue smears using A O or Giemsa stain as described in
Section 1.
REFERENCES
1.
A L D A Y - S A N Z V., R O D G E R H., T U R N B U L L T., A D A M S A. & RICHARDS R . H.
( 1 9 9 4 ) . - An immunohistochemical diagnostic test for rickettsial disease.
J. Fish Dis., 17, 1 8 9 - 1 9 1 .
2.
BRANSON E.J. & D I A Z - M U N O Z D . N .
( 1 9 9 1 ) . - Description of a new disease
condition occurring in fanned coho salmon, Oncorhynchus
(Walbaum), in South America. J. Fish Dis., 14, 1 4 7 - 1 5 6 .
3.
4.
BRAVO S. ( 1 9 9 4 ) . - First report of Piscirickettsia salmonis in freshwater.
American Fisheries Society Fish Health Section Newsletter, 22(1), 6 .
BRAVO S.S. & CAMPOS M . L . ( 1 9 8 9 ) . - Coho salmon syndrome in Chile.
American Fisheries Society Fish Health Section Newsletter,
5.
kisutch
17(2), 3 .
BROCKLEBANK J . R . , S P E A R E D . J . , A R M S T R O N G R . D . & EVELYN T. ( 1 9 9 2 ) . -
Septicaemia suspected to be caused by a rickettsia-like agent in farmed
Atlantic salmon. Can. Vet. J., 33, 4 0 7 - 4 0 8 .
6.
CVTTANICH J . D . , G A R A T E N . O . & C E . SMITH ( 1 9 9 0 ) . - Etiological agent in
Chilean coho disease isolated and confirmed by Koch's postulates.
American Fisheries Society Fish Health Section Newsletter, 18(1), 1-2.
7.
CVITANICH J . D . , G A R A T E N . O . & SMITH C E . ( 1 9 9 1 ) . - The isolation of a
rickettsia-like
organism causing disease and mortality in Chilean
salmonids and its confirmation by Koch's postulates. J. Fish Dis., 14, 1 2 1 145.
8.
DAVIES A.J. ( 1 9 8 6 ) . - A rickettsia-like organism from dragonets,
Callionymus lycra L . (Teleostei: Callionymidae) in Wales. Bulletin of the
European Association of Fish Pathologists, 6(4), 1 0 3 .
Piscirickettsiosis
139
9.
EVELYN T.P.T. ( 1 9 9 2 ) . - Salmonid rickettsial septicemia. In: Kent M L . ,
ed. Diseases of Seawater Netpenreared Salmonid Fishes in the Pacific
Northwest. Canadian Special Publication of Fisheries and Aquatic
Sciences No. 1 1 6 . Dept. Fisheries and Oceans, Nanaimo, Canada, 1 8 - 1 9 .
10.
FRYER J . L . & LANNAN C.N. ( 1 9 9 0 ) . - Isolation of a Rickettsiales-like
organism from diseased Coho salmon {Oncorhynchus kisutch) in Chile.
Proceedings, Annual Meeting of the Japanese Society of Fish Pathology,
March 2 9 - 3 1 , Tokyo, Japan. 2 6 pp.
11.
FRYER J . L . & LANNAN C.N. ( 1 9 9 0 ) . - Preliminary characterization of a
rickettsia-like pathogen of salmonids (Abstract). - In: Vlllth International
Congress of Virology. International Union of Microbiological Societies,
Berlin, 4 9 .
12.
FRYER J . L . & LANNAN C.N. ( 1 9 9 4 ) . - Rickettsial and chlamydial infections.
In: Stevenson R.M. & Woo P.T.K., eds. Fish Diseases II: Bacterial and
Fungal Infections. Commonwealth Agriculture Bureau International,
Wallingford. In press.
13.
FRYER J . L . , LANNAN C.N.,
GARCES L . H . , LARENAS J.J.
& SMITH
P.A.
( 1 9 9 4 ) - Piscirickettsia salmonis, a newly described intracellular pathogen
of salmonid fish. In: Program and Abstracts of the Second International
Marine Biotechnology Conference. Society for Industrial Microbiology,
Arlington, Virginia, 5 6 7 - 5 7 2 .
14.
FRYER J.L., LANNAN C.N.,
GARCES L . H . , LARENAS J.J. & SMITH
P.A.
( 1 9 9 0 ) . - Isolation of a rickettsiales-like organism from diseased coho
salmon {Oncorhynchus kisutch) in Chile. Fish Pathology, 25, 1 0 7 - 1 1 4 .
15.
F R Y E R J.L., LANNAN C.N.,
GIOCANNONI S.J.
& W O O D N.D.
(1992).
-
Piscirickettsia salmonis gen. nov., sp. nov., the causative agent of an
epizootic disease in salmonid fishes. Int. J. Systemic Bacterio!, 42, 1 2 0 126.
16.
GARCES L . H . , LARENAS J.J.,
SMITH P.A.,
SANDINO S., LANNON C.N.
&
FRYER J.L. ( 1 9 9 1 ) . - Infectivity of a rickettsia isolated from coho salmon,
Oncorhynchus kisutch. Dis. Aquat. Org., 11, 9 3 - 9 7 .
17.
HOSKINS G ( 1 9 9 2 ) . - A report on Paranthesis disease in British Columbia.
Pacific Biological Station, Nanaimo, BC, Canada, l p .
18.
LANNAN C.N. ( 1 9 9 4 ) - Extracellular survival of Piscirickettsia
Fish Dis., In press.
salmonis. J.
140
19.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
LANNAN C . N . , CVITANICH J . D . & EVELYN T.P.T. (1992). - Rickettsial
Disease of Salmonids. Pacific Northwest
Committee Informational Report 6. 11 pp.
20.
Fish
Health
Protection
LANNAN C.N., EWING S.A. & FRYER J . L . (1991). - A fluorescent antibody
test for detection of the rickettsia causing disease in Chilean salmonids. J.
Aquatic Animal Health, 3 , 229-234.
21.
LANNAN C.N. & FRYER J . L . (1991). - Recommend methods for inspection
of fish for the salmonid rickettsia. Bulletin of the European Association of
Fish Pathologists, 11, 135-136.
22.
LANNAN C.N. & FRYER J . L . (1993). - Piscirickettsia salmonis, a major
pathogen of salmonid fish in Chile. Fisheries Research, 17, 115-121.
23.
L A U E R B . A . , R E L L E R L . B . & M I R R E T T S. (1981). - Comparison of acridine
orange and Gram stains for detection of microorganisms in cerebrospinal
fluid and other clinical specimens. J. Clin. Microbiol., 14, 201-205.
24.
M O H A M E D Z. (1939). - The discovery of a rickettsia in a fish. Minist.
Agrie. Cairo, Tech. Sci. Serv., Vet. Sect. Bull. 214, 6 pp.
25.
O L S E N A.B.,
EVENSEN O.,
SPEILBERG L . , M E L B Y H.P.
(1993). - "Ny" Iaksesykdom forârsaket av rickettsie. Norsk
12, 40-41.
&
HASTEIN
T.
Fiskeoppdrett.,
26.
O Z E L M. & SCHWANZ-PFITZNER I. (1975). - Vergleichende elektronenmikroskopische Untersuchungen an Rhabdoviren pflanzlicher und
tierischer Herkunft: III. Egtved-Virus (VHS) der Regenbogenforelle
(Salmo gairdneri) und Rickettsienahnliche Organismen. Zbl. Bakt. Hyg. I.
AbtOrig., 230, 1-14.
27.
RODGER H.D. & DRINAN E.M. (1993). - Observation of a rickettsia-like
organism in Atlantic salmon. Salmo salar L . , in Ireland. J. Fish Dis., 16,
361-369.
28.
SCHÂFER J . W . , ALVARADO V., ENRIQUES R. & M O N R A S M. (1990). - The
"coho salmon syndrome" (CSS): a new disease in Chilean salmon, reared
in sea water. Bulletin of the European Association of Fish
Pathologists,
10, 130.
29.
W O L F K. (1981). - Chlamydia and rickettsia of fish. Fish Health
10(3), 1-5.
News,
Pathogens of molluscs
141
DISEASES OF BIVALVE MOLLUSCS
D I S E A S E S N O T I F I A B L E T O T H E OIE
C H A P T E R 15
DIAGNOSTIC TECHNIQUES: GENERAL INFORMATION
1.
PRELIMINARY REMARKS
Diagnostic procedures for molluscan pathogens are limited, and
histological examination is the conventional method applicable to these
hosts. Consequently we can present in a single document all the techniques
currently available for the diagnosis of diseases believed to be contagious.
This presentation takes into account the performance of these techniques
in terms of sensitivity, ease of use, and samples to be examined.
2.
SAMPLING
2.1
Sampling points
For each zone referred to, a number of sampling points must be
selected so as to maximise the chances of detecting pathogens. Account
must be taken of parameters having an effect on the development of the
pathogenic agents, such as stocking density, water flows, and the
development cycle of the molluscs.
For a given zone at least three sampling points must be selected. The
number of points must be increased for large zones containing several
discrete areas of cultivation of the susceptible species.
Whenever possible, at least one sample must be taken from natural
beds. Any molluscs showing abnormalities (abnormal growth, gaping
shells) must be selected.
2.2
Sampling period and frequency
The frequency of inspection is based on the infection period, and
inspections have to take place thereafter. Inspection periods must also
take account of the transfer of molluscs, which generally takes place in
spring and autumn. Sampling should therefore be carried out:
•
once a year after the summer period (i.e. in September-October in
the northern hemisphere) for the genera Marteilia,
Haplospohdia
and Perkinsus;
•
twice a year (spring/autumn) for the genera Bonamia
Mikrocytos and for iridovirus.
and
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OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
2.3
Sampling size
During the initial two-year period which precedes the granting of
approved status, the sample size for each sampling point is 150 or a
sufficient number to ensure detection at a 9 5 % confidence level of
pathogen carriers at a prevalence of 2%.
During the subsequent years (maintenance of approved status), the
sample size must be maintained at 150 to ensure detection at a 9 5 %
confidence level of pathogen carriers at a prevalence of 2%.
3.
SHIPMENT O F SAMPLES
All molluscs sampled must be delivered to the approved laboratory within
24 h after sampling. They must be packed in accordance with current
standards in order to keep them in good condition. A label clearly stating
the place of sampling and the health history (if any) must be attached to
the sample.
4.
M A C R O S C O P I C EXAMINATION
The molluscs must be opened carefully so as not to damage the tissues, in
particular the mantle, gills, heart and digestive gland. Anomalies and
lesions of the tissues, as well as any shell deformities, shell-boring
organisms and conspicuous mantle inhabitants, must be noted.
5.
EXAMINATION O F STOCKS WHERE ABNORMAL MORTALITIES OCCUR
Whenever abnormal mortalities occur in stocks of bivalve molluscs an
urgent investigation must be carried out to determine the aetiology. (An
abnormal mortality is a sudden sizeable mortality [more than 15% of the
stock] which occurs in a short time between two observations [15 days]. In
the hatchery an abnormal mortality is a failure of successive productions of
larvae coming from different broodstock.)
The sample taken must consist of 150 individual oysters and must be
handled in accordance with the procedure defined for histological analysis.
This technique must be used initially, before any other type of
examination. The samples are fixed, preferably in Carson's fixative, which
allows re-use of the sample for electron microscopy.
Bonamiosis
143
CHAPTER
16
BONAMIOSIS
(B431)
SUMMARY
Bonamiosis is a lethal infection of the blood cells of common
oysters, accompanied by nonspecific branchial lesions. It is caused
by two protozoa of the phylum Ascetospora, Bonamia ostreae and
Bonamia sp.
The disease may occur throughout
experimentally by cohabitation or
the year, and can be
inoculation.
transmitted
Bonamiosis affects the following oysters: Ostrea edulis, O. angasi,
O. conchaphila and Tiostrea chilensis. Its geographical
distribution
is: Spain, USA, France, Greece, Irish Republic, Italy, Netherlands,
UK, New Zealand and Australia.
In the natural environment, diagnosis is possible after an oyster has
spent 3-4 months in an infected area, and it is accomplished by
histological techniques in sections or smears of infected organs.
For diagnosis, the rules for sampling are those stated in the general
section (Chapter 15).
DIAGNOSTIC PROCEDURES
1.
P R E P A R A T I O N AND E X A M I N A T I O N O F S A M P L E S F O R
1.1
BONAMIA
Cytological examination: blood smears
For larvae and oysters, after the samples have been air-dried, make a
squash for larvae or put the cardiac tissues on a histological slide. The
slides are air-dried, then fixed in methanol. The prepared larvae and
oysters are stained using a commercially available blood staining kit, in
accordance with the manufacturer's instructions. After staining they are
rinsed using tap water and allowed to dry completely with cold or
warm air, and mounted in a synthetic resin.
The parasite (2 pm in size) shows up as blue (cytoplasm) with a red
nucleus. It may be observed inside or outside the haematocytes. An
observation time of 5 min per slide is sufficient.
1.2
Histology
Cut the bivalve through the digestive gland, gills and heart and place
the sample in a fixative liquid such as Davidson's, Bouin's or Carson's
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
144
fluid. Carson's fixative enables the samples to be used for electron
microscopy when necessary. The sample volume:fixative volume ratio
of 1 : 1 0 must be respected.
Several non-specific stains, e.g. haematoxylin-eosin or Masson's
trichrome process, enable Bonamia to be visualised. These examples
are not exhaustive. It is recommended that two sections per oyster be
examined.
The parasite ( 2 um in size) occurs freely in the connective tissue and
haematocytes. Detection of microcells in haematocytes, connective
tissue or extracellularly should be confirmed by transmission electron
microscopy.
REFERENCES
1.
BUCKE D., HEPPER B., K E Y D. & BANNISTER C.A. ( 1 9 8 4 ) . - A report on
Bonamia ostreae in Ostrea edulis in the UK. Cons. Inter. Explor. Mer,
CM 1984/K, 9, 1-7.
2.
DINAMANI
P . , HINE
P.M.
&
JONES
J.B.
(1987).
-
Occurrence
and
characteristics of the haemocyte parasite Bonamia sp. in the New Zealand
dredge oyster Tiostrea lutaria. Dis. Aquat. Org., 3, 3 7 - 4 4 .
3.
E L S T O N R.A., FARLEY C.A. & K E N T M.L. ( 1 9 8 6 ) .
- Occurrence
and
significance of bonamiasis in European flat oysters Ostrea edulis in North
America. Dis. Aquat. Org., 2, 4 9 - 5 4 .
4.
5.
G R I Z E L H . ( 1 9 8 5 ) . - Etudes des récentes épizooties de l'huitre plate Ostrea
edulis L. et de leur impact sut l'ostréiculture bretonne. Thèse de doctorat,
Université des Sciences et Techniques de Languedoc, Montpellier, France.
G R I Z E L H . , C O M P S M., RAGUENNES D., LEBORGNE Y . , T I G E G. & M A R T I N
A.G. ( 1 9 8 3 ) . - Bilan des essais d'acclimation â'Ostrea chilensis
côtes de Bretagne. Rev. Trav. Inst. PêchesMarit., 46, 2 0 9 - 2 2 5 .
sur les
6.
HINE P . M . ( 1 9 9 1 ) . - The annual pattern of infection by Bonamia sp. in
New Zealand flat oysters, Tiostrea chilensis. Aquaculture, 93, 2 4 1 - 2 5 1 .
7.
M C A R D L E J . F . , M C K I E R N A N F., F O L E Y H . & JONES D . H . ( 1 9 9 1 ) . - The
current status of Bonamia disease in Ireland. Aquaculture,
8.
M O N T E S J. & M E L E N D E Z I. ( 1 9 8 7 ) . - Données sur la parasitose de Bonamia
ostreae
chez l'huitre plate de Galice, côte Nord-Ouest de l'Espagne.
Aquaculture,
9.
93, 2 7 3 - 2 7 8 .
67, 1 9 5 - 1 9 8 .
P I C H O T Y . , C O M P S M., T I G E G , GRIZEL H . & RABOUIN M.A. ( 1 9 7 9 ) . -
Recherches sur Bonamia ostreae gen. n., sp. n., parasite nouveau de
l'huitre plate Ostrea edulis L. Rev. Trav. Inst. Pêches Marit., 43, 1 3 1 - 1 4 0 .
Bonamiosis
10.
145
T I G E G . , G R I Z E L H . , COCHENNEC N. & RABOUIN M . A . ( 1 9 8 4 ) . - Evolution
de la situation épizootiologique en Bretagne en 1 9 8 3 suite au
développement de Bonamia ostreae. Cons. Inter. Explor. Mer, CM
1984/F, 14, 1 - 1 0 .
11.
VAN BANNING P . ( 1 9 8 2 ) . - Some aspects of the occurrence, importance and
control of the oyster pathogen Bonamia ostreae in the Dutch oyster
culture. Proc. Inter. Coll. Invertebr. Pathol., 6 - 1 0 Sep. 1 9 8 2 , 2 6 1 - 2 6 3 .
12
V A N BANNING P . ( 1 9 8 7 ) . - Further results of the Bonamia
challenge tests in Dutch oyster culture. Aquaculture, 6 7 , 1 9 1 - 1 9 4 .
ostreae
147
Haplosporidiosis
C H A P T E R 17
HAPLOSPORIDIOSIS
(B432)
SUMMARY
Haplosporidiosis
is an infection, sometimes lethal, of blood cells,
connective
tissue and digestive tissues, often accompanied
by
brownish-red discolouration
of gills and mantle. It is caused by
protozoa of the phylum Ascetospora:
Haplosporidium nelsoni, H.
costale, H. armoricanum and Haplosporidium spp. The following
species are affected: Crassostrea virginica, Ostrea edulis, Ruditapes
decussatus, and Ostrea angasi. Only H. nelsoni, a parasite of C.
virginica, is responsible for epizootics. This species is found in the
bays of Delaware and Chesapeake (eastern coast of USA). It is also
reported from Florida to Massachusetts and in Maine.
Infection takes place between mid-May and the end of October. It
has not been possible to transmit the disease experimentally in the
laboratory.
Haplosporidia
may be detected in smears or stained sections of
infected organs. For diagnosis, the rules for sampling are those
defined in the general section (Chapter 15).
DIAGNOSTIC PROCEDURES
1.
P R E P A R A T I O N AND E X A M I N A T I O N O F S A M P L E S F O R
1.1
HAPLOSPORIDIA
Cytological examination
Cut the digestive gland and the gills along a sagittal plane, soak up
excess water by applying absorbent paper to the sample, then press that
part of the cut surface with the digestive gland, gills and mantle against
a glass slide. The slides are dried in air and then fixed with methanol
(for 2-3 min).
The slides are stained using a commercially available staining kit in
accordance with the manufacturer's instructions. Dip the slides in the
first bath for 4-5 seconds, then immerse immediately in the second bath
(3 s e c ) . Rinse with tap water, dry completely in cold or warm air, and
mount in synthetic resin (Eukitt). Plasmodium stages (4 to 30 pm in
size) show up as blue cytoplasm with red nuclei. They affect mainly the
gills, palps, connective tissue and epithelium of the digestive gland.
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
148
1.2. Histological examination
Cut the visceral mass along a sagittal plane with small scissors, and
place the sample in a fixative (Davidson's, Bouin's or Carson's fluid);
the last-named is suitable for samples which may be examined later by
electron microscopy if necessary. There should be at least 1 0 parts of
fluid to every part of sample by volume.
The sections are subsequently treated by conventional histological
procedures. Many nonspecific stains show up Ascetospora: haemalumeosin, Masson's trichrome and others. Two sections from each oyster
should be examined. The different stages of the parasite can be
observed in the gills, palps, connective tissue and epithelium of the
digestive gland.
REFERENCES
1.
ANDREWS J.D. & W O O D J.L. ( 1 9 6 7 ) . - Oyster mortality studies in Virginia
- VI. History and distribution of Minchinia
in Virginia. Chesapeake Science, 8, 1 - 1 3 .
2.
nelsoni, a pathogen of oyster,
BURRESON E.M., R O B I N S O N M.E. & VILLALBA A. ( 1 9 8 8 ) . - A comparison
of paraffin histology and hemolymph analysis for the diagnosis of
Haplosporidium
nelsoni (MSX) in Crassostrea virginica (Gmelin). J.
Shellfish Res., 7 , 1 9 - 2 3 .
3.
COUCH J.A. & ROSENFIELD A. ( 1 9 6 8 ) .
- Epizootiology
of
Minchinia
costalis and Minchinia nelsoni in oysters introduced into Chintoteague
Bay, Virginia. Proc. Natl. Shellfish Assoc., 58, 5 1 - 5 9 .
4.
5.
FARLEY C.A. ( 1 9 6 7 ) . A proposed life cycle of Minchinia
nelsoni
(Haplosporida, Haplosporidiidae) in the American oyster
Crassostrea
virginica. J. Protozool., 14, 6 1 6 - 6 2 5 .
F O R D S.E., FIGUERAS L.A.J. & HASKIN H.H. ( 1 9 9 0 ) . - Influence of selective
breeding, geographic origin, and disease on gametogenesis and sex ratios
of oysters, Crassostrea virginica, exposed to the parasite Haplosporidium
nelsoni (MSX). Aquaculture, 88, 2 8 5 - 3 0 1 .
6.
FORD
S.E.
& HASKIN H.H.
Haplosporidium
nelsoni
1 9 5 7 - 8 0 . J. Invertebr.
7.
(1982).
- History
and
epizootiology
of
(MSX), an oyster pathogen in Delaware Bay,
Pathol,
40, 1 1 8 - 1 4 1 .
HASKIN H.H. & F O R D S.E. ( 1 9 8 2 ) . - Haplosporidium
nelsoni
(MSX) on
Delaware Bay seed oyster beds: a host-parasite relationship along a salinity
gradient. J. Invertebr. Pathol, 40, 3 8 8 - 4 0 5 .
Haplosporidiosis
8.
PERKINS F.O. ( 1 9 6 8 ) . - Fine structure of the oyster pathogen
nelsoni (Haplosporida, Haplosporidiidae). J. Invertebr. Pathol,
149
Minchinia
10, 2 8 7 -
307.
9.
PERKINS F.O. ( 1 9 6 9 ) . - Electron microscope studies of sporulation in the
oyster pathogen Minchinia
costalis (Sporozoa: Haplosporida). - J.
Parásito!.,
55, 8 9 7 - 9 2 0 .
151
Marteiliosis
CHAPTER
18
MARTEILIOSIS
(B434)
SUMMARY
Marteiliosis
is caused by protozoan parasites
of the phylum
Ascetospora, genus Marteilia, which develop mainly in the epithelial
cells of the digestive gland and are associated with emaciation of the
oyster and exhaustion of its reserves of energy (glycogen).
The type species, Marteilia refringens, is a lethal parasite of the
common European oyster, Ostrea edulis. Marteiliosis affects mainly
the following
species of shellfish:
Ostrea edulis, Saccostrea
commercialis, Mytilus edulis, Mytilus galloprovincialis, Saccostrea
cucullata, Crassostrea virginica and Cardium edule. Diseases of
economic significance are caused by Marteilia refringens in Ostrea
edulis, and Marteilia sydneyi in S. commercialis.
The geographical distribution of these diseases is: Australia, Spain,
France, Italy, Morocco. The period of infection for M. refringens is
confined to the summer, when water temperature is greater than
17°C. M. sydneyi infections occur all year round. The mode of
infection and the life cycle outside the host are unknown.
Diagnosis is by examination of smears of infected organs, or stained
sections. For diagnosis, the rules for sampling are those stated in the
general section (Chapter 15).
DIAGNOSTIC PROCEDURES
1.
P R E P A R A T I O N AND E X A M I N A T I O N O F S A M P L E S F O R
1.1
MARTEILIA
Cytological examination
In order to prepare the smears, cut a section through the digestive
gland and the gills, remove the excess water by placing the sample on
blotting paper, then place on a slide the sample corresponding to the
section which passes through the digestive tract The slides are dried in
air and then fixed in methanol (2-3 min).
The samples are stained using a commercially available staining kit.
Stain in accordance with the manufacturer's instructions. After
staining, rinse using tap water and allow to dry completely with cold or
warm air and place in a synthetic resin.
152
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
The parasite is 5-8 um in size in the early stages and may reach up to
40 um during sporulation. The cytoplasm of the cells stains a blue
colour of a greater or lesser intensity, the nucleus being bright red. The
secondary cells or sporoblasts are surrounded by a bright halo. An
observation time of 5 min per slide is sufficient.
1.2
Histological examination
For histological sections, cut a section through the digestive gland
using small scissors, and place the sample in a fixative liquid such as
Davidson's, Bouin's or Carson's. The last-named enables the samples to
be reused for electron microscopy if necessary. The ratio of volume of
tissue to volume of fixative must be no more than 1:10.
The samples are subsequently handled in accordance with the classical
histological methods. Several non-specific stains allow
Marteilia
refringens to be observed: haemalum-eosin, Masson's trichrome. These
examples are not exhaustive. It is recommended that two sections per
oyster be examined.
The young stages of Marteilia are present in the epithelium of the
stomach; later developed stages can be found in the epithelium of the
digestive diverticulata. Free sporangiae can also be observed in the
lumen of the intestine.
REFERENCES
1.
C O M P S M., GRIZEL H. & PAPAYANNI Y. (1982). - Infection parasitaire
causée par Marteilia
maurini
sp. n. chez la moule
galloprovincialis.
Cons. Inter. Explor. Mer, CM, F: 24, 2 pp.
2.
Mytilus
G R I Z E L H., C O M P S M., BONAMI J.R., COUSSERANS F., D U T H O I T J.L. & L E
PENNEC M.A. (1974). - Recherche sur l'agent de la maladie de la glande
digestive de Ostrea edulis L. Science et Pêche, 240, 7-30.
3.
G R I Z E L H., C O M P S M., COUSSERANS F., B O N A M I J.R. & V A G O C. (1974). -
Etude d'un parasite de la glande digestive observée au cours de l'épizootie
actuelle de l'huitre plate. C. R. Acad. Sci. Paris, 279, D, 783-784.
4.
GRIZEL H. & TIGE G . (1979). - Observations sur le cycle de
refringens. Haliotis, 8, 327-330.
5.
M O Y E R M.A., BLAKE N.J. & ARNOLD W.S. (1993). - An
disease causing mass mortality in the Atlantic calico scallop,
gibbus (Linnaeus, 1758). J. Shellfish Res., 12, 305-310.
Marteilia
acetosporan
Argopecten
Marteiliosis
6.
7.
153
PERKINS F.O. (1976). - Ultrastructure of sporulation in the European flat
oyster pathogen Marteilia
refringens.
Taxonomic implication. J.
Protozool., 23, 64-74.
ROUBAL F.R., M A S E L J. & LESTER R.J.G. (1989). - Studies on
sydneyi, agent of QX disease in the Sydney rock oyster,
commercialis, with implications for its life cycle. Aust. J. Mar.
Res., 40, 155-167.
Marteilia
Saccostrea
Freshwater
8.
PERKINS F.O. & W O L F P.H. (1976). - Fine structure of Marteilia sydneyi
sp. n. - Haplosporidian pathogen of Australian oysters. J. Parasitoi, 62,
528-538.
9.
TIGE G. & RABOUIN M A . (1976). - Etude d'un lot de moules transferees
dans un centre touché par l'épizootie affectant l'huître plate. Cons. Inter.
Explor. Mer, CM, K: 18, 1-7.
10.
W O L F P . H . (1972). - Occurrence of a Haplosporidian in Sydney Rock
oysters (Crassostrea
commercialis)
from Morton Bay, Queensland,
Australia. J. Invertebr. Pathol., 19, 416-417.
11.
W O L F P.H. (1979). - Life cycle and ecology of Marteilia sydneyi in the
Austalian oyster, Crassostrea commercialis. Mar. Fish Rev., 41, 70-72.
155
Mikrocytosis
C H A P T E R 19
MIKROCYTOSIS
(B436)
SUMMARY
Mikrocytosis is a lethal disease of two genera of oysters, Crassostrea
gigas and Saccostrea commercialis, caused by two protozoa (Protista
incerta sedis), Mikrocytos mackini and M. roughleyi respectively.
The disease induces pustules, abscesses and ulcers, mainly on the
mantle, with corresponding brown scars on the shell. Abscesses are
composed of granular haemocytes and hyalinocytes which contain
small cells of 1-3 ¡xm. The mortality rate varies around 40%. The
disease occurs more often during the spring for C. gigas and during
the austral winter for S. commercialis.
The geographical distribution of M. mackini is the west coast of
Canada, probably ubiquitous throughout the strait of Georgia
including Henry Bay, Denman Island and confined to other specific
localities around Vancouver Island. M. roughleyi occurs in New
South Wales, Australia.
Diagnosis is by examination of smears of infected organs or stained
sections. For diagnosis, the rules for sampling are those stated in the
general section (Chapter 15).
DIAGNOSTIC
1.
PROCEDURES
P R E P A R A T I O N AND E X A M I N A T I O N O F S A M P L E S F O R M I K R O C Y T O S I S
1.1
Cytological examination
Cut a section through the abscesses or ulcers, remove the excess water
by placing the sample on blotting paper, then blot on a slide the sample
corresponding to the section which passes through the infected tissue.
The slides are dried in air and then fixed with methanol (2-3 min).
The slides are stained using any equivalent Wright-Giemsa stain (e.g.
Merck's Hemacolor Kit or Baxter's Diff-Quick) in accordance with the
manufacturer's instructions. Dip the slides in the first bath for 4-5
seconds, then immerse immediately in the second bath (3 sec). Rinse
with tap water, dry completely in cold or warm air, and mount in
synthetic resin (Eukitt).
OIE Diagnostic Manual for Aquatic Animal Diseases, 1 9 9 5
156
The parasite, 1-3 pm in diameter, appears included in haemocytes or
free of the host cells and has blue cytoplasm and a small red nucleus.
An observation time of 5 min per slide is sufficient.
1.2
Histological examination
For histological sections, cut a section through the body of the oyster
including pustules, abscesses and ulcers if any are present. Then place
the sample in a fixative liquid such as Davidson's, Bouin's or Carson's.
The last-named enables the samples to be reused for electron
microscopy if necessary. T h e ratio of volume of tissue to volume of
fixative must be no more than 1:10.
The samples are subsequently handled in accordance with the classical
histological methods. Several non-specific stains allow Mikrocytos to
be observed: haemalum-eosin, Masson's trichrome. These examples are
not exhaustive. It is recommended that two sections per oyster be
examined.
The stages of Mikrocytos are present in the haemocytes concentrated in
and around the abscesses or the ulcers for M. roughleyi and in the
vesicular connective tissue cells on the periphery of the lesions for M.
mackini.
REFERENCES
1.
BOWER S.M. ( 1 9 8 8 ) . - Circumvention of mortalities caused by Denman
Island oyster disease during mariculture of Pacific oysters. Amer. Fish.
Soc. Special Publication, 18, 2 4 6 - 2 4 8 .
2.
FARLEY C.A., W O L F P.H. & E L S T O N R A . ( 1 9 8 8 ) . - A long-term study of
'microceir disease with a description of a new genus, Mikrocytos (g. n.),
and two new species, Mikrocytos
mackini (sp. n.) and Mikrocytos
roughleyi (sp. n.). Fishery Bulletin, 86, 5 8 1 - 5 9 3 .
Perkinsosis
157
C H A P T E R 20
PERKINSOSIS
(B433)
SUMMARY
Perkinsosis is a lethal infection of connective tissue, caused by
protozoan parasites of the phylum Apicomplexa, genus Perkinsus
(formerly Dermocystidium). The type species, Perkinsus marinus, is
a lethal parasite of the American oyster, Crassostrea virginica.
The geographical
distribution
Perkinsus species is: Australia,
Coast).
of the disease caused by various
Spain, Portugal,and the USA (East
Principally
the following
shellfish
are affected:
Crassostrea
virginica, Ruditapes decussatus, R. philippinarum, Haliotis rubra, H.
laevigata, Tridacna gigas and recently Argopecten irradians.
Diseases of economic importance are caused by Perkinsus marinus
in C. virginica (USA), Perkinsus atlanticus in R. decussatus
(Portugal), and Perkinsus olseni in Haliotis rubra (Australia) and H.
laevigata Some 50 species of bivalves may harbour Perkinsus
species without harmful effect.
The main period of infection is summer, when the water temperature
exceeds 20°C. The infective stage is a biflagellate zoospore, which
develops into a trophozoite. Trophozoites multiply within the host by
successive binary fissions.
Perkinsus is detectable in stained sections and by culture in
thioglycollate medium. For diagnosis, the rules for sampling are
those stated in the general section (Chapter 15).
DIAGNOSTIC PROCEDURES
1.
P R E P A R A T I O N AND E X A M I N A T I O N O F SAMPLES F O R
1.1
PERKINSUS
Histological examination
Cut the visceral mass with small scissors along a sagittal plane, and
place the sample in a fixative (Davidson's, Bouin's or Carson's fluid);
the last-named is suitable for samples which may be examined later by
electron microscopy if necessary. There should be at least ten parts of
fluid to every part of sample by volume.
Samples are then treated by conventional histological procedures.
158
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Many nonspecific stains show up Apicomplexa: haemalum-eosin,
Masson's trichrome and others. Two sections from each oyster should
be examined.
According to species, the size of trophozoites varies from 5-7 pm for P.
marinus, 13-16 pm for P. olseni and 10-15 pm for P. atlánticas.
Trophozoites are usually characterized by the presence of a vacuole
which displaces the nucleus towards the periphery. With haemalumeosin stain, the cytoplasm of trophozoites stains pink and the nucleus
violet.
In the case of Ruditapes decussatus, the presence of trophozoites elicits
a cystic reaction, and the cysts may be large enough to be seen with the
naked eye.
1.2. Diagnosis by culture in thioglycollate medium
Tissue samples measuring 5 x 1 0 m m are cut out (giving preference to
rectal samples from oysters) and placed in liquid thioglycollate medium
(Difco) plus antibiotics. The antibiotics (benzylpenicillin 0.5 mg/ml
and streptomycin sulphate 0.5 mg/ml) are added under sterile
conditions after having been sterilised by filtration (0.22 pm).
Incubation is done at 22-25°C for 48-72 hours.
The size of cultured parasites increases from 3-10 to 70-125 pm. After
incubation, the fragments of tissues are collected and placed in Lugol's
1:5 solution for 10 min, then mounted between a slide and a cover slip
for microscopic examination in the fresh state. Perkinsus parasites are
present as round cells, stained blue or bluish-black by Lugol's solution,
and they may reach 120 pm.
REFERENCES
1.
ANDREWS J.D. (1965). - Infection experiments in nature with
Dermocystidium marinum in Chesapeake Bay. Chesapeake Science, 6, 6067.
2.
ANDREWS J.D. (1988). - Epizootiology of the disease caused by the oyster
pathogen Perkinsus marinus and its effects on the oyster industry. Amer.
Fish. Soc. Special Publication, 18,47-63.
3.
AZEVEDO C. (1989). - Fine structure of Perkinsus atlanticus n. sp.
(Apicomplexa, Perkinsea) parasite of clams, Ruditapes decussatus, from
Portugal. J. Parasitoi, 75, 627-635.
4.
LESTER R.J.G. & DAVIS G.H.G. (1981). - A new Perkinsus species
(Apicomplexa, Perkinsea) from the abalone, Haliotis ruber. J. Invertebr.
Pathol., 37, 181-187.
Perkinsosis
5.
LESTER R . J . G . ,
159
GOGGIN C L .
&
SEWELL K . B . ( 1 9 9 0 ) .
- Perkinsus
in
Australia. In: Cheng T.C. & Perkins F.O., eds. Pathology in marine
science. Academic Press, New York.
6.
MACKIN J . G . ( 1 9 5 1 ) . - Histopathology of infection of Crassostrea virginica
gmelin by Dermocystidium
marinum Mackin, Owen and Collier. Bull.
Marine Science of the Gulf and Caribbean, 1, 7 2 - 8 7 .
7.
MACKIN J . G . ( 1 9 6 2 ) . - Oyster disease caused by Dermocystidium
marinum
and other micro-organisms in Louisiana. Publ. Inst. Mar. Sci., 1 3 2 - 2 2 9 .
8.
MCGLADDERY
S.E.,
CAWTHORN
R.J.
&
BRADFORD
B.C.
Perkinsus karlssoni n. sp. (Apicomplexa) in bay scallops
irradians. Dis. Aquat. Org., 10, 1 2 7 - 1 3 7 .
(1991).
-
Argopecten
9.
PERKINS F.O. ( 1 9 8 8 ) . - Structure of protistan parasites found in bivalve
molluscs. Amer. Fish. Soc. Special Publication, 18, 9 3 - 1 1 1 .
10.
PERKINS F.O. & M E N Z E L R . W . ( 1 9 6 7 ) . - Ultrastructure of sporulation in
the oyster pathogen Dermocystidium
marinum.
J. Invertebr.
Pathol., 9,
205-229.
11.
R A Y S.M. ( 1 9 5 2 ) . - A culture technique for the diagnosis of infections with
Dermocystidium marinum Mackin, Owen and Collier in oysters. Science,
116,
12.
360-361.
R A Y S.M. ( 1 9 6 6 ) . - A review of the culture method of detecting
Dermocystidium marinum with suggested modifications and precautions.
Proc. Natl. Shellfish Assoc., 54, 5 5 - 6 9 .
161
Iridoviroses (B431)
CHAPTER 21
IRIDOVIROSES
(B435)
SUMMARY
Three iridovirus infections have been reported in bivalve molluscs.
Although the viruses have not yet been purified and studied in detail,
they have been classified as iridovirus because of the presence of
DNA and the shape and size of their virions, 380 nm in diameter. The
first cases of infection described were in the Portuguese
oyster,
Crassostrea angulata, in which the virus elicited necrosis
of
branchial tissues and infections of haemocytes. Virions develop from
inclusion bodies, present in infected cells.
Iridoviruses can affect both C. angulata and C. gigas (larvae). Lethal
iridoviral infections in C. angulata produce lesions of the gills, or
lesions of haemocytes and connective tissues. In C. gigas the
infection is particularly responsible for the death of larvae, and it is
known as oyster velar virus disease (OWD).
The geographical distribution of these diseases, which are no longer
being reported, was Spain, France and Portugal. Larval mortality
has affected hatcheries on the western coast of USA.
In the absence of established cell lines from bivalve molluscs, the
only diagnostic procedure is to examine stained smears or sections
of infected organs, supported in presumptive cases by electron
microscopy. More sensitive and easier methods are needed for
routine diagnosis.
For diagnosis, the rules for sampling are those stated in the general
section (Chapter 15).
DIAGNOSTIC PROCEDURES
1.
P R E P A R A T I O N AND E X A M I N A T I O N O F SAMPLES F O R
1.1
IRIDOVIRUS
Cytological examination
Cut the digestive gland and gills along a sagittal plane, soak up excess
water by applying absorbent paper, then press that part of the cut
sample which passes through affected organs against a glass slide. The
slides are dried in air and then fixed with methanol (2-3 min).
162
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
The slides are stained using Merck's Hemacolor Kit (with reagent
solution 2 [ref. 11956] for red staining and reagent solution 3 [ref.
11957] for blue staining). Dip the slides in the first bath for 4-5
seconds, then immerse immediately in the second bath (3 sec). Rinse
with tap water, dry completely in cold or warm air, and mount in
synthetic resin (Eukitt).
It is sufficient to examine each slide for 5 min.
The cytoplasm of infected cells stains blue, and it contains a weaklystaining red nucleus and an inclusion body, variable in size, stained
bright red.
1.2. Histological examination
Cut the visceral mass and the gills with small scissors along a sagittal
plane, and place the sample in a fixative (Davidson's, Bouin's or
Carson's fluid); the last-named is suitable for samples which may be
examined later by electron microscopy if necessary. There should be at
least ten parts of fluid to every part of sample by volume.
The samples are then treated by conventional histological procedures.
Many nonspecific stains show up iridoviral inclusion bodies:
haemalum-eosin, Masson's trichrome and others. Two sections from
each oyster should be examined.
Infected cells contain a highly chromophilic inclusion body and are
larger than normal.
1.3. Electron microscopy
With EM, the viruses described during the outbreaks of gill disease and
the massive mortalities of 1970 were present in large polymorphic cells
of 30-40 um and in conjunctive tissue cells respectively. These cells
contained inclusion bodies from which extruding viral particles could
be observed. The virions were icosahedral and they were 380 nm and
350 nm in diameter respectively (according to the disease), with
electron-dense nucleoids of 250 nm and 190 nm.
The iridovirus of oyster velar velum disease infects only the epithelial
cells of the velum. It is also icosahedral and is 230 nm in diameter.
Iridoviroses ( B 4 3 1 )
163
REFERENCES
1.
COMPS M . ( 1 9 7 0 ) . - La maladie des branchies chez les huîtres du genre
Crassostrea, caractéristiques et évolution des altérations, processus de
cicatrisation. Rev. Trav. Inst. Pêches Mari t., 34, 2 3 - 4 4 .
2.
COMPS M. ( 1 9 7 2 ) . - Observations sur la résistance d'huîtres du genre
Crassostrea au cours de la mortalité massive de 1 9 7 0 - 1 9 7 1 dans le bassin
de Marennes-Oléron. Cons. Inter. Explor. Mer, CM. K 22, 9 pp.
3.
COMPS M. & DUTHOIT J.L. ( 1 9 7 6 ) . - Infection virale associée à la maladie
des branchies de l'huître portugaise Crassostrea
Acad. Sci. Paris, 283, D, 1 5 9 5 - 1 5 9 6 .
4.
angulata
LmK. C. R.
C O M P S M., B O N A M I J.R., V A G O C. & CAMPILLO A. ( 1 9 7 6 ) . - Une viróse de
l'huître portugaise (Crassostrea
angulata LmK).
C. R. Acad. Sci. Paris,
282, D, 1 9 9 1 - 1 9 9 3 .
5.
COMPS M. & M A S S O R.M. ( 1 9 7 8 ) . - Study with fluorescent technique of
the virus infection of the Portuguese oyster Crassostrea angulata LmK.
Proc. Inter. Coll. Invertebr. Pathol., 1 1 - 1 7 Sept. 1 9 7 8 , 3 9 - 4 0 .
6.
COMPS M. ( 1 9 8 3 ) . - Recherches histologiques et cytologiques sur les
infestions intracellulaires des mollusques bivalves marins. Thèse Doct.
Etat Sci. Nat., Montpellier, 1 2 8 pp.
7.
COMPS M. ( 1 9 8 8 ) . - Epizootic diseases of oysters associated with viral
infections. Amer. Fish. Soc. Special Publication, 1 8 , 2 3 - 3 7 .
8.
ELSTON R. ( 1 9 7 9 ) . - Virus-like particles associated with lesions in larvae
of Pacific oysters (Crassostrea gigas). J. Invertebr. Pathol., 33, 7 1 - 7 4 .
9.
ELSTON R. ( 1 9 8 0 ) . - Ultrastructural aspects of a serious disease of hatchery
reared larval oysters, Crassostrea gigas Thünberg. J. Fish Dis., 3, 1 - 1 0 .
10.
ELSTON R.A. & WILKINSON M.T. ( 1 9 8 5 ) . - Pathology, management and
diagnosis of oyster velar virus disease ( O W D ) . Aquaculture,
48, 1 8 9 - 2 1 0 .
Baculoviral midgut gland necrosis virus
165
DISEASES OF CRUSTACEANS
OTHER SIGNIFICANT DISEASES
C H A P T E R 22
BACULOVIRAL MIDGUT GLAND NECROSIS VIRUS
(B443)
SUMMARY
Baculoviral midgut gland necrosis virus (BMN) has been found in
cultured kuruma shrimp, Penaeus japonicus, in Japan and Korea. In
Japan it is considered one of the major problems in hatcheries. As
with Baculovirus penaei (BP) and Penaeus monodon-(y/?e baculovirus
(MBV), the main target organ for BMN is also the hepatopancreas,
and the virus may initiate mass mortality for the infected larvae.
However, unlike BM and MBV, it does not produce an occlusion
body in the infected
hepatopancreatocyte.
So far BMN epizootics have been reported in Japan. BMN-like virus
(non-occluded, type C baculovirus) has been reported from Penaeus
monodon in the Philippines, Australia and Indonesia.
Experiments on artificial infection revealed that BMN caused high
mortality
to healthy
P. japonicus larvae. BMN has
been
experimentally
transmitted to P. monodon, P. chinensis and P.
semisulcatus.
Two diagnostic techniques, wet-mount and histopathology
of the
hepatopancreas,
have been used for the demonstration of BMN
infection in P. japonicus. Under electron microscopy,
enveloped
virions measuring approximately 310 nm x 72 nm were observed. No
occlusion bodies have been found in the infected nuclei of
hepatopancreatocytes.
Since the oral route has been demonstrated to be the main infection
pathway for BMN infection, complete or partial eradication of viral
infection may be accomplished by thorough washing of fertile eggs
or nauplii using clean sea water.
A. I N T R O D U C T I O N
Epizootics of baculoviral midgut gland necrosis (BMN) of larval Penaeus
japonicus have occurred in the Kyushu and Chugoku area of Japan since 1971.
BMN-like virus (non-occluded, type C baculovirus) has been reported from P.
monodon in the Philippines and possibly in Australia and Indonesia. BMN virus
166
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
was experimentally
semisulcatus.
transmitted
to P.
monodon,
P.
chinensis
and
P.
During recent years, P. japonicus has become one of the most important
cultured shrimps in Taiwan; BMN has not been detected in that species or in
other cultured species in that country (unpublished finding).
The B M N infected moribund shrimp larvae showed a nuclear hypertrophy and
remarkable cellular necrosis or collapse of the midgut gland (hepatopancreas).
Results of infectivity trials and field surveys showed that BMN may initiate
mass mortality for the healthy larvae. It was also noted that, as for MBV and
BP, viruses released with faeces into the environmental water of intensive
culture systems of P. japonicus play an important role in disease spread.
B. D I A G N O S T I C T E C H N I Q U E S
The morbid or heavily infected larvae reveal the clinical sign of a cloudy midgut
gland, which may easily be observed by the naked eye. In addition, wet-mounts
and histopathology of hepatopancreas are more reliable techniques for a definite
diagnosis of BMN virus infection.
1.
W E T - M O U N T TECHNIQUE
Infection of BMN virus can be rapidly diagnosed by demonstration of fresh
squashes of hepatopancreas under dark field or light microscopy. In
hepatopancreas of diseased post larvae with no polyhedral or round
occlusion bodies, but with hypertrophied nuclei in midgut gland epithelial
cells as viewed under dark field illumination equipped with a wet type
condenser, infected nuclei appear white against the dark background due
to the increased reflected and diffracted rays produced by numerous virus
particles in the nucleus. Feulgen stain makes the difference clearer
between normal nuclei (about 10 pm in diameter) and infected
hypertrophied nuclei (about 20-30 pm in diameter). The healthy
hepatopancreas only presents very few fat droplets. A similar result may
also be obtained when 10% formalin is used for fixation.
2.
HlSTOPATHOLOGICAL TECHNIQUE
Shrimps for observation were fixed in neutral 10% formalin and Bouin's
solution, stained with hematoxylin and eosin by Vago-Amargier's method
and observed under conventional microscopy. The M B V infected shrimps
showed greatly hypertrophied nuclei within hepatopancreatic epithelial
cells that are undergoing necrosis. The abnormal nuclei reveal marginated
chromatin, a laterally disassociated nucleolus, and the absence of occlusion
bodies that characterise infections by type A and type B baculoviruses.
Baculoviral midgut gland necrosis virus
167
C. ERADICATION PROCEDURES
Studies on the virucidal effect of disinfectants on BMN virus demonstrated that
this virus was inactivated by 5 ppm of chlorine, 2 5 ppm of iodine, 1 0 0 ppm of
benzalkonium chloride or benzethonium chloride and 0 . 5 % of formalin.
However, these solutions at each described concentration level were toxic to
shrimp larvae. Therefore, no disinfectants can be used in practice in hatcheries
for eradication of BMN virus infection. It is suggested that washing nauplii or
fertile eggs with clean sea water to eradicate the digested excrement of shrimp
is a better way for the production of BMN virus-free larvae. Suggested
procedures for eradication of BMN virus from hatcheries are shown in Table 1.
Table 1
Suggested procedures for eradication of B M N infection
Broodstock
zz>
=>
Collection of
Pass through a soft gauze with
fertile eggs
—> pore size of 8 0 0 um to remove
for 1 minute
digested excrement or faeces of
shrimp
Wash with running sea water at salinity level of 2 8 - 3 0 % for 3 - 5 minutes
to make sure all the faecal debris have been removed.
Pass through a soft gauze with pore size of 1 0 0 um to collect eggs or
nauplii.
Wash with running sea water at salinity level of 2 8 - 3 0 % for 3 - 5 minutes
to remove the adhesive viral particles.
Hatchery pond.
REFERENCES
1.
LIGHTNER D.V. ( 1 9 8 3 ) . - Diseases of cultured Penaeid shrimp. In: J.P.
McVey (éd.). CRC Handbook of Mariculture. Vol. 1, Crustacean
Aquaculture. CRC Press. Boca Raton, Florida, USA, pp. 2 8 9 - 3 2 0 .
2.
LIGHTNER D.V. ( 1 9 8 8 ) . - Baculoviral midgut gland necrosis (BMN)
disease of Penaeus japonicus. In: C.J. Sindermann and D.V. Lightner
(eds). Disease Diagnosis and Control in North American Marine
Aquaculture. Elsevier, Amsterdam, 2 6 - 2 9 .
3.
LIGHTNER D.V. & REDMAN R.M. ( 1 9 8 9 ) . - Hosts, geographic range and
diagnostic procedures for the penaeid virus diseases of concern to shrimp
culturists in Americas. In: P. De Loach, W.J. Dougherty & M A . Davidson
(eds). Frontiers of Shrimp Research. Elsevier, Amsterdam, 1 7 3 - 1 9 6 .
168
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
4.
M O M O Y A M A K. (1989). - Virucidal effect of some disinfectants on
baculoviral midgut gland necrosis (BMN) virus. Fish Pathology, 24(1),
47-49.
5.
NATIVIDAD J.M. & LIGHTNER D.V. (1992). - Prevalence and geographic
distribution of MBV and other diseases in cultured giant tiger prawns
(Penaeus monodori) in the Philippines. In: W. Fulks and K . L . Main (eds).
Diseases of Cultured Penaeid Shrimp in Asia and the United States. The
Oceanic Institute, Honolulu, Hawaii, 139-160.
6.
PARK M.A. (1992). - The status of culture and diseases of penaeid shrimp
in Korea. In: W. Fulks and K . L . Main (eds). Diseases of Cultured Penaeid
Shrimp in Asia and the United States. The Oceanic Institute, Honolulu,
Hawaii, 161-167.
7.
SANO T. & M O M O Y A M A K. (1992). - Baculovirus infection of penaeid
shrimp in Japane. In: W. Fulks and K . L . Main (eds). Diseases of Cultured
Penaeid Shrimp in Asia and the United States. The Oceanic Institute,
Honolulu, Hawaii, 169-174.
8.
SANO T.,
NISHIMURA T.,
FUKUDA H.,
HAYASHIDA T.
& M O M O Y A M A K.
(1984). - Baculoviral midgut gland necrosis (BMN) of kuruma shrimp
{Penaeus japonicus)
larvae in Japanese intensive culture systems.
Helgolander Meeresunters, 37, 255-264.
9.
SANO T.,
NISHIMURA T.,
FUKUDA H.,
HAYASHIDA T.
& M O M O Y A M A K.
(1985). - Baculoviral infectivity trials on kuruma shrimp (Penaeus
japonicus), of different ages. In: Fish and Shellfish Pathology. Academic
Press, New York, pp. 397-403.
10.
SANO T., NISHIMURA T., O G U M A K . , M O N O Y A M A K . & T A K E N O N . (1981).
- Baculovirus infection of cultured Kuruma shrimp Penaeus japonicus
Japan. Fish Pathology, I S , 185-191.
11.
in
V A G O C. & AMARGIER A. (1963). - Coloration histologique pour la
différenciation des corps d'inclusion polyhèdriques de virus d'insectes.
Ann. Epiphyties, 14(3), 269-274.
Nuclear polyhedrosis baculoviroses
169
C H A P T E R 23
NUCLEAR POLYHEDROSIS BACULOVIROSES
{Penaeus monodon-type
baculovirus and
Baculoviruspenaei)
(B441 / B 4 4 2 )
SUMMARY
Penaeus monodon-type baculovirus (MBV) and Baculovirus penaei
(BP), two nucelar polyhedrosis viruses, have been considered to be
potentially serious pathogens in the larval stages of host shrimp.
They possess a wide distribution and diverse host range. Both MBV
and BP are characterised by the presence of occlusion bodies (OBs),
which may be referred as polyhedral occlusion bodies (POBs), or
polyhedral inclusion bodies (PIBs), in hepatopancreatocytes,
gut
cells and digested excrement.
Studies on the epizootiology of MBV infection in P. monodon showed
a very high incidence of MBV in postlarvae, juvenile and broodstock
of P. monodon obtained from Asia and Southeastern Asia. P.
monodon from Texas, Ecuador and Brazil also revealed
positive
MBV infection. The other species revealed a low MBV incidence rate
in these areas. BP was also demonstrated to be widespread in
distribution in cultured and wild penaeid shrimps in the Northern
and Southern Americas.
Crowding and chemical or environmental stress may enhance the
pathogenicity
and increase the prevalence of these two viruses in
their hosts.
MBV can be diagnosed
by wet-mount and
histopathological
observation of hepatopancreas.
Observation of digested
excrement
for the presence of MBV occlusion bodies is also feasible.
Prevention of MBV and BP infection in hatcheries may be obtained
by first washing nauplii or fertile eggs with formalin,
iodophores
and clean sea water.
A. I N T R O D U C T I O N
Baculoviruses of shrimps including Penaeus monodon-type baculovirus (MBV)
and Baculovirus penaei (BP), two nuclear polyhedrosis viruses, have been
considered to be potentially serious pathogens in the larval stages of host
shrimp. They possess a wide distribution and diverse host range. Both MBV
and BP are characterised by the presence of occlusion bodies (OBs), which may
be referrred to as polyhedral occlusion bodies (POBs) or polyhedral inclusion
bodies (PIBs), in hepatopancreatocytes, gut cells and digested excrement.
Penaeus monodon-typz
baculovirus (MBV) is found in shrimps from the
170
OIE Diagnostic Manual for Aquatic Animal Diseases: Prototype, 1995
Indopacific and Pacific Coasts of Asia, Australia, Africa, Southern Europe, and
Northern and Southern America.
M B V has been found to infect various species of shrimps including Penaeus
esculentus, P. kerathurus, P. merguiensis,
P. monodon, P. plebejus,
P.
penicillatus, P. semisulcatus, P. vannamei, and Metapenaeus ensis. The most
serious infection is found in the cultured black tiger prawn, P. monodon.
Epizootiological studies on MBV infection in P. monodon in 1989 showed an
M B V incidence of more than 5 0 % in postlarval juveniles and broodstocks
obtained from Taiwan and Southeastern Asia (unpublished result). BP was
found in cultured and wild penaeid shrimps in the Americas, ranging from the
Northern Gulf of Mexico south through the Caribbean and reaching at least as
far as central Brazil, and from Peru to Mexico along the Pacific Coast. No BP
has ever been found in wild or cultured penaeids in Asia. This virus was
demonstrated to infect Penaeus aztecus, P. brasiliensis, P. duorarum, P.
marginatus,
P. paulensis,
P. penicillatus,
P. schmitti,
P. setiferus,
P.
stylirostris, P. subtilis, and P. vannamei.
Experiments performed in hatcheries showed that M B V initiated mortality for
infected postlarval P. monodon. Results obtained from pathogenicity studies
showed that environmental stress significantly affects the survival of MBVinfected P. monodon and P. penicillatus. Mass mortality which occurred in
MBV-infected P. monodon postlarvae may have been initiated by mixed
infection with Vibrio spp. However, when the shrimps were kept in a grow-out
pond with good environmental conditions, no mortality or growth retardation
occurred in the MBV-infected juveniles. Eradication of M B V infection in P.
monodon or P. penicillatus
in hatcheries, which will be described in the
following section, is therefore important.
It was also demonstrated that BP is pathogenic to P. vannamei, P. aztecus,
P. duorarum, and P. marginatus. As for MBV, physical or chemical stress on
BP-infected shrimps may enhance the severity of virus infection and increase
the prevalence rate.
B. D I A G N O S T I C T E C H N I Q U E S
Several diagnostic procedures are used in screening shrimp stocks for MBV
infections:
1)
2)
3)
4
Wet-mount examination of hepatopancreas
Histopathological observation of hepatopancreas
Examination of excrement
Recombinant DNA technique.
The first two techniques are employed for studies on the epizootiology and
pathogenicity of M B V and BP. However, the third technique is suitable for
screening of MBV and BP-free broodstocks for hatcheries. The fourth technique
provides a highly specific and sensitive method for detection of M B V and BP in
Nuclear polyhedrosis baculoviroses
171
shrimp stocks. Furthermore, this technique can be used to identify the target
organs of the viral infection and to determine which life stages are most
susceptible.
1.
W E T - M O U N T TECHNIQUE
M B V infection may readily be diagnosed by the demonstration of spherical,
round, pyramidal or tetrahedral shaped occlusion bodies in wet-mounts of
hepatopancreatocytes or midgut of shrimp under bright field or phase
contrast microscopy. The occlusion bodies may also be pyramidal or
tetrahedral. BP infected hepatopancreatocytes reveal tetrahedral occlusion
bodies. However, the sensitivity of this techniques is rather limited since
spherical or tetrahedral occlusions are difficult to distinguish from lipid
droplets and secretory granules in unstained wet-mounts of tissue squashes.
Aqueous malachite green (0.05%) may be used in preparing for MBV and
BP diagnosis to aid in the observation of occlusion bodies. Also, occlusion
bodies (in squashes or histology) fluoresce under ultraviolet light following
staining with aqueous 0.18% phloxine, thereby providing a rapid and
specific diagnosis.
2.
HISTOPATHOLOGICAL TECHNIQUE
This technique is a reliable technique for definite diagnosis of MBV and BP
infection. Prior to preparation for observations, the specimen should be
preserved in Davidson's fixative containing 3 3 % ethyl alcohol (95%), 2 0 %
formalin (approximately 3 7 % formaldehyde), 11.5% glacial acetic acid and
33.5% distilled or tap water.
In addition, Bouin's fixative, Carnoy's solution and 10% formalin solution
may also be used for this purpose. To obtain a better result, no dead shrimps
should be used. Live, moribund or compromised shrimps are killed by
injection of fixative, then the cuticle over the céphalothorax and abdomen
just lateral to the dorsal midline are opened and immersed in fixative for 2472 hrs, then transferred to 7 0 % ethyl alcohol for storage. These fixed
specimens may be shipped by wrapping in cloth or paper towel saturated
with 7 0 % ethyl alcohol and packed in plastic bags. Tissues such as the
hepatopancreas or gills may be cut into small pieces and fixed directly in the
fixative.
After dehydration, the specimens are embedded in paraffin and sectioned
into 5-7 um thickness. Harris hematoxylin and eosin, Giemsa's and Gram's
staining solutions may be used for the demonstration of spherical (MBV) or
tetrahedral (BP) occlusion bodies in hepatopancreatocytes, gut epithelial
cells or gut lumen.
Necrosis and loss of hepatopancreatic and midgut epithelial cells are
observed in the sections of diseased shrimps. Single or more often multiple
occlusion bodies in hypertrophied nuclei, and chromatin diminution and
margination in infected hepatopancreatic epithelial cells, have been observed
in advanced infection. Occlusion bodies may be stained bright red with
172
OIE Diagnostic Manual for Aquatic Animal Diseases: Prototype, 1995
haematoxylin and eosin, and stained intensely with Gram's stain and
toluidine blue. Brown and Benn histological Gram stain, although not
specific for baculovirus occlusion bodies, tends to stain occlusions more
intensely (either red or purple, depending on section thickness, time of
decolonisation, etc.) than the surrounding tissue, aiding in demonstrating
their presence in low-grade infections.
3.
DIGESTED EXCREMENT EXAMINING TECHNIQUE
The shrimps used for examination may be placed in an aquarium or plastic
tank for a few hours until digested excrement is present. The faeces are
collected and placed in a glass centrifuge tube. Subsequently, digesting
solution consisting of 4 parts n-butanol, 1 part n-hexane and 10 parts
distilled water is added to each tube and the mixture is then homogenised for
3-5 minutes and centrifuged at a speed of 1,500-2,000 rpm. Similar
procedures may be repeated twice for better digestion. The sediments are
observed for the presence of MBV or BP occlusion bodies with the aid of
staining using eosin, Giemsa, acridine orange or malachite green solutions.
4.
RECOMBINANT D N A TECHNIQUE
The recombinant D N A technique involves probe cloning, probe labelling,
filter hybridisation analysis, and in situ hybridisation analysis. Standard
procedures are involved in these components of the recombinant DNA
technique, and they can be found in most molecular biology manuals. Many
of the reagents needed for the procedures can even be purchased as kits. The
DNA of both MBV and BP has been successfully detected through the
application of gene probes. Details of the methods can be found in the
literature. General descriptions of the procedures are provided below.
4.1
Probe cloning and labelling
BP or MBV virions can be isolated from hepatopancreatic tissue of infected
shrimp; they are then purified by centrifugation. The DNA can also be
purified with phenol extraction and ethanol precipitation. The BP or M B V
DNA is digested with restriction endonucleases, ligated into the bacterial
vector, and used to transform bacterial cells. Clones containing viral DNA
inserts are selected. Gene probes can be labelled either radioactively with P
or S or with dioxigenin, a non-radioactive label. Convenient kits containing
reagents for labelling are available from Amersham, DuPont-NFN,
Promega, BRL, and other companies.
4.2
Filter hybridisation analysis
Shrimp to be tested are processed for DNA extraction and purification.
The DNA is blotted onto a nylon filter and subsequently the filter is
prehybridised, then hybridised with a labelled BP or M B V probe.
Hybridisation mixtures will usually include formamide (30-50%),
sheared fragments of DNA, dextran sulphate, Denhardt's solution, and
NaCl in sterile water. After hybridisation, the filter is washed several
173
Nuclear polyhedrosis baculoviroses
times in salt solution to wash off non-specifically bound probe. The
hybridisation signal can be detected using suitably sensitive
autoradiography film if a radiolabeled probe is used. Detection of the
non-radiolabelled probe is usually through an enzyme reporter, such as
alkaline phosphatase or peroxidase; a colour precipitate can then be
observed at the hybridisation site.
4.3
In situ hybridisation
This procedure is now well established for the identification and
localisation of BP and MBV DNA in tissue sections of infected shrimp.
Before attempting in situ hybridisation, it is usual to fix the viral DNA
in place in the cell with a fixative such as formalin. The tissue is then
washed in PBS, embedded in paraffin, and sectioned at approximately
4-6 pm thickness. Sections can be collected onto precoated slides and
then deparaffinised. The sections can then be rehydrated. This is
followed by digestion with proteinase K (10 pg/ml) for 30 min at 37°C.
Subsequently the sections are prehybridised for 1 hour at 37°C, and
then hybridised by adding labelled MBV or BP DNA probes. After
hybridisation, the sections can be washed and the presence of viral
DNA can be detected within the cells.
Table 1
Suggested procedures for the production of BP or M B V
non-infected postlarvae
a) Nauplii*
Collection of
nauplii
using plankton net
^>
^>
^>
Formalin 400 ppm
for 30 seconds
to 1 minute
-z>
Hatchery ponds
Running
sea water
for 1-2 minutes
-^>
Formalin
100 ppm
for 1 minute
Running
sea water
for 3-5 minutes
-^
Hatchery ponds
Running
sea water
for 1-2 minutes
Running
sea water
for 3-5 minutes
Iodophore
0.1 ppm
for 1 minute
:
b) Fertilised eggs**
Collection of
fertilised eggs
Iodophore
0.1 ppm
for 1 minute
^>
Nauplii are much easier to collect than are fertilised eggs in hatcheries
Fertilised eggs are more sensitive than nauplii to formalin.
174
5.
OIE Diagnostic Manual for Aquatic Animal Diseases: Prototype, 1995
ERADICATION PROCEDURES
Experiments performed on the infection mechanism showed that M B V
infection was initiated by the oral route. Eradication of MBV infection in
hatcheries may therefore be obtained by several washes of nauplii or
fertilised eggs using clean sea water, formalin and iodophore solutions
(Table 1). Nauplii are much easier to collect than are fertilised egg in
hatcheries, and fertilised eggs are more sensitive to formalin. The
procedures used for nauplii are therefore highly recommended for
commercial hatcheries.
REFERENCES
1.
A N D E R S O N I.G., SHARIFF M., N A S H G. & N A S H M . (1987). - Mortalities of
juvenile shrimp Penaeus monodon, associated with Penaeus
monodon
baculovirus, cytoplasmic reo-like virus and rickettsial and bacterial
infections, from Malaysian brackish water ponds. Asian Fisheries Science,
1, 47-64.
2.
BROCK J.A., L I G H T N E R D . V . & B E L L T . A . (1983). - A review of four virus
(BP, MBV, B M N and IHHN) diseases of penaeid shrimp with particular
references to clinical sgnificance, diagnosis and control in shrimp
aquaculture. International Council for the Exploration of the Sea. C M .
1983/Gen: 10/Mini-Symposium.
3.
B R U C E L . D . , REDMAN R . M . & LIGHTNER D . V . (1994). - Application of
gene probes to determine target organs of a penaeid shrimp baculovirus
using in situ hybridisation. Aquaculture, 1 2 0 , 45-51.
4.
B R U C E L . D . , R E D M A N R . M . , L I G H T N E R D . V . & B O N A M I J . R . (1993). -
Application of gene probes to detect a penaeid shrimp baculovirus in fixed
tissue using in situ hybridisation. Dis. Aquatic Organisms, 1 7 , 215-221.
5.
6.
CHEN S.M., L o C F . , Liu S.M. & K o u G.H. (1989). - T h e first
identification of Penaeus monodon baculovirus (MBV) in cultured sand
shrimp, Metapenaeus ensis. Bull. Eur. Assoc. Fish Pathologists, 9(3), 6 2 64.
C H E N S.N., C H A N G P.S., K o u G.H. & LIGHTNER D . V . (1989). - Studies on
virogenesis and cytopathology of Penaeus monodon baculovirus (MBV) in
giant tiger prawn (Penaeus monodon) and the red tail prawn (Penaeus
penicillatus). Fish Pathology, 2 4 , 89-100.
7.
CHEN
S.N., C H A N G P.S. & K o u G.H. (1989).
-
Observation
on
pathogenicity and epizootiology of Penaeus monodon baculovirus (MBV)
in cultured shrimps in Taiwan. Fish Pathology, 2 4 , 189-195.
Nuclear polyhedrosis baculoviroses
8.
175
CHEN S.N., C H A N G P . S . & K o u G . H . ( 1 9 9 4 ) . - Rapid and histopathological
diagnosis of Penaeus monodon baculovirus infection in shrimp. Report on
Fish Disease Research, 16 (in press).
9.
C H E N S.N.,
C H A N G P.S. & K o u G.H.
(1990).
- Infection
route
and
eradication of Penaeus monodon baculovirus (MBV) in larval giant tiger
prawns, Penaeus monodon. In: W . Fulks & K.L. Main (eds). Diseases of
Cultured Penaeid Shrimp in Asia and the United States. Oceanic Institute,
Honolulu, Hawaii, USA, 1 7 7 - 1 8 4 .
10.
COUCH J.A. ( 1 9 7 4 ) . - An enzootic nuclear polyhedrosis virus of pink
shrimp: ultrastructure, prevalence, and enhancement. J. Inv. Pathol, 24,
311-331.
11.
LESTER
R.J.G.,
DOUBROVSKY
A.,
PAYNTER
J.L.,
SAMBHI
S.K.
&
ATHERTON J.G. ( 1 9 8 7 ) . - Light and electron microscope evidence of
baculovirus infection in the prawn Penaeus plebejus. Dis. Aquatic
Organisms,
12.
3(2), 217-219.
L I G H T N E R D . V . & R E D M A N R . M . ( 1 9 8 1 ) . - A baculovirus-caused disease of
the penaeid shrimp, Penaeus monodon. J. Inv. Pathol., 38. 2 9 9 - 3 0 2 .
13.
LIGHTNER D . V ( 1 9 8 3 ) . - Diseases of cultured Penaeid shrimp. In: J.P.
McVey (éd.). CRC Handbook of Mariculture. Vol. 1, Crustacean
Aquaculture. CRC Press. Boca Raton, Florida, USA, pp. 2 8 9 - 3 2 0 .
14.
LIGHTNER D. V , R E D M A N R.M. & B E L L T.A. ( 1 9 8 3 ) . - Observations on the
geographic distribution, pathogenesis and morphology of the baculovirus
from Penaeus monodon Fabricius. Aquaculture, 3 2 ( 3 / 4 ) , 2 0 9 - 2 3 3 .
15.
LIGHTNER D.V., REDMAN R.M., WILLIAMS R.R., M O H N E Y L.L., CLERX
J.P.M., B E L L T.Z. & B R O C K J.A. ( 1 9 8 5 ) . - Recent advances in penaeid
virus disease investigations. J. World Mariculture
16.
Society, 16, 2 6 7 - 2 7 4 .
LIGHTNER D . V , HEDRICK R.P., F R Y E R J.L., C H E N S.N., L I A O I.C. & K o u
G.H. ( 1 9 8 7 ) . - A survey of cultured penaeid shrimp in Taiwan for viral
and other important diseases. Fish Pathology, 22(3), 1 2 7 - 1 4 0 .
17.
L I G H T N E R D . V . ( 1 9 8 8 ) . - Diseases of cultured penaeid shrimp and prawns.
In: C.J. Sindermann & D.V. Lightner (eds). Disease Diagnosis and
Control in North American Marine Aquaculture. Elsevier, Amsterdam, pp.
8-127.
18.
LIGHTNER D . V ,
REDMAN
R.M.
&
ALMADA
RUIZ
E.A.
(1989).
-
Baculovirus penaei in Penaeus stylirostris (Crustacea: Decapoda) cultured
in Mexico: unique cytopathology and new geographic record. J. Inv.
Pathol.,
53, 1 3 7 - 1 3 9 .
176
19.
OIE Diagnostic Manual for Aquatic Animal Diseases: Prototype, 1995
L u C.C., T A N G F . J . K , K O U G.H. & C H E N S.N. (1993). - Development of a
Penaeus monodon-type baculovirus (MBV) DNA probe by polymerase
chain reaction and sequence analysis. J. Fish Dis., 16, 551-559.
20.
Lu C C ,
T A N G F . J . K , K O U G.H. & C H E N S.N. (1995). - Detection of
Penaeus monodon-type baculovirus (MBV) infection in Penaeus
Fabricius by in situ hybridisation. (Manuscript in preparation.)
monodon
21.
L U N A L . G . (ed). (1968). - Manual of Histological Staining Methods of the
Armed Forces Institute of Pathology. McGraw-Hill Book Company, New
York.
20.
T H U R M A N R.B.,
L I G H T N E R D.V.
&
HAZANOW
S.
(1989).
-
Unique
physicochemical properties of the occluded penaeid shrimp baculoviruses
and their use in the diagnosis of infections. J. World Aquaculture Soc, 20,
75A-76A (abstract).
Infectious hypodermal & haematopoietic necrosis virus
177
C H A P T E R 24
INFECTIOUS HYPODERMAL & HAEMATOPOIETIC
NECROSIS VIRUS
(B444)
SUMMARY
Infectious hypodermal and haematopoietic necrosis (IHHN) virus, a
parvo- or a picorna-like
virus, is considered to be a potential
pathogen for a serious disease in several species of shrimps. This
virus invades all the cells in tissues of ectodermal and mesodermal
origin. However, in light cases, the endoderm-derived tissues such as
midgut, midgut caeca and hepatopancreas are not infected by this
virus.
IHHN is characterised by its worldwide distribution among cultured
shrimp. However, no data are available on the distribution of this
virus in wild shrimp. The geographic distribution of IHHN virus
infection in shrimp has increased very significantly in recent years.
Infection by IHHN virus has been demonstrated to cause disease in
Penaeus stylirostris, P. monodon and P. vannamei.
Wet-mount
or histopathological
observations
are the
main
techniques for demonstration of IHHN virus infection in shrimps. For
quarantine purposes, bioassay of a suspect shrimp population with a
sensitive
indicator
species
followed
by
wet-mount
or
histopathological
observation
of randomly
selected
indicator
shrimps is recommended. To enhance the infection, shrimps may be
reared under relatively crowded and stressful conditions.
Monoclonal antibodies against IHHNV may be applied to detect
IHHNV infected shrimps with a very high specificity by using
indirect
enzyme-linked
immunosorbent
assay
(ELISA),
and
immunoblot and Western blot assays. Also, the in situ hybridisation
technique can be applied on paraffin sections of IHHNV
infected
shrimps using a DNA probe.
A. I N T R O D U C T I O N
Infectious hypodermal and haematopoietic necrosis (IHHN) virus, a parvo- or a
picorna-like virus, has been found in shrimps cultured on the Atlantic and
Pacific coasts, Pacific Islands, Asia and Middle East. The virus invades all the
cells in tissues of ectodermal and mesodermal origin.
This virus is also characterised by its wide host range. Penaeus vannamei,
P.
178
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
stylirostris, P. monodon and P. semisulcatus are susceptible to IHHN virus in
nature. However, experimental infection of IHHN virus was obtained in P.
schmitti, P. japonicus, P. aztecus and P. duorarum. Infection by IHHN virus
was demonstrated to cause acute epizootics and mass mortality for P.
stylirostris, P. monodon and P. vannamei. In contrast no significant mortality
was found in other penaeid shrimps. IHHN virus initiated higher mortality rates
(up to 80 or 90%) in populations of small size. Although infection of tissue cells
were observed in older or larger shrimps, very little mortality was obtained.
It is concluded that like BP or MBV, IHHN virus affects the larvae or young
juvenile stages of shrimp more seriously than it affects larger shrimps.
No eradication technique has been developed. However, IHHNV free larvae may
be produced by using virus free broodstock.
B.
DIAGNOSTIC
TECHNIQUES
The following three diagnostic procedures are employed to diagnose IHHN
virus infection in shrimps: a) direct samplings for histopathological
observations; b) enhancement of infection followed by direct sampling
techniques', c) bioassay of a suspect population with a sensitivity 'indicator'
species.
1.
D I R E C T EXAMINATION
Samples for examining are fixed in Davidson's fixative and stained by
haematoxylin and eosin staining procedures for histopathological
observations as described in the chapter on MBV and BP (Appendix 2)
under bright field microscopy with 200 or 400x magnification. Cowdry
type A eosinophilic inclusion bodies are observed in hypertrophic nuclei of
the tissues or organs including gills, pereiopods, maxillipeds, stomach
mucosa, gnathorax, ventral nerve cord and ganglia, and the
haematopoietic tissues. Necrosis and inflammatory response of infected
tissue cells is the other sign for IHHN virus infection which may result in
shrimp mortality.
2.
ENHANCEMENT OF INFECTION
The enhancement procedure may be achieved with a quarantined
population by rearing shrimps in relatively crowded or stressful conditions.
The stressors may include O2 deficiency in water, unsuitable temperature,
insufficient food supply or presence of N H 3 or NO2 in the water. All these
stressful conditions may initiate a heavy infection of IHHN virus in the
host shrimps. The moving of larvae from a hatchery pond to an aquarium
or tank would also act as a stressor to the infected shrimps.
Infectious hypodermal & haematopoietic necrosis virus
3.
179
BlOASSAY T E C H N I Q U E
Carriers of IHHN virus may be detected with highly sensitive 'clean'
(uninfected) shrimps such as juvenile P. stylirostris at body weight of 0.054 g. This technique is considered to be the best method for examination or
screening of IHHNV infected shrimps or carriers.
The following two methods can be applied to expose the suspect carrier
shrimps to indicator species:
1)
Injection or immersion of 'indicator' shrimps with homogenates
derived from suspected samples. Using this technique 'indicator'
shrimps will show signs of EHHN virus disease 5-20 days after
treatment.
2)
Rearing suspect carrier shrimps with indicator shrimps. Using this
method 'indicator' shrimps will show signs of IHHN disease within 1
or 2 months after rearing.
In addition, chopped carcases of suspect 'carrier' shrimp may also be fed to
'indicator' shrimps. Fifteen to 60 days following feeding, the indicator
shrimps will reveal signs of IHHN disease.
4.
M O N O C L O N A L ANTIBODIES T O I H H N V
IHHNV virus may be purified from infected shrimps. Crude homogenate
from infected shrimps may be clarified by a series of low-speed
centrifugations and the virus pelleted at 145,000 g. The virus suspension is
treated with activated charcoal, filtered on a Celite-235 bed and extracted
with freon. The virus is repelleted at 145,000 g and further purified on a
15-40% sucrose gradient followed by a 25-40% caesium chloride gradient.
Monoclonal antibodies (MAbs) may be obtained by routine techniques
using myeloma cells and spleen cells of BALB/C strain mice. MAbs may
react specifically with IHHNV in immunoblots and Western blots.
5.
G E N E PROBES
IHHNV gene probes may be obtained by purification of virus and
extraction of DNA followed by transformation of DNA in competent E.
coli DH5 cells. Positive reactions may be obtained when the probe is tested
on paraffin sections of IHHNV infected shrimps. Tested shrimps may be
fixed in Davidson's fixative, embedded and sectioned at 5 um using
standard histological procedures. IHHNV probes react only with IHHNV
infected cells.
REFERENCES
1.
B E L L T.A. & LIGHTNER D.V. (1988). - A Handbook of Normal Shrimp
Histology. Special Publication No. 1, World Aquaculture Society, Baton
Rouge, Louisiana, USA. 114 pp.
180
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
2.
B E L L T A . & LIGHTNER D.V. (1987). - IHHN disease of Penaeus
stylirostris: effects of shrimp size on disease expression. J. Fish Diseases,
10, 165-170.
3.
B O N A M I J.R.,
BREHELIN M.,
M A R I J., TRUMPER B.
& LIGHTNER
D.V.
(1990). - Purification and characterisation of IHHN virus of penaeid
shrimps. J. Gen. Virol, 7 1 , 2657-2664.
4.
BROCK J.A., L I G H T N E R D . V . & B E L L T . A . (1983). - A review of four virus
diseases of penaeid shrimp with particular references to clinical
significance, diagnosis and control in shrimp aquaculture. International
Council for the Exploration of the Sea. C M . 1983/Gen.: 10/MiniSymposium.
5.
LIGHTNER D.V. (1983). - Diseases of cultured Penaeid shrimp. In: J.P.
McVey (ed.). CRC Handbook of Mariculture. Vol. 1, Crustacean
Aquaculture. CRC Press, Boca Raton, Florida, USA, pp. 289-320.
6.
LIGHTNER
D.V.,
REDMAN
R.M.
&
BELL
TA.
(1983).
-
Infectious
hypodermal and hematopoietic necrosis (IHHN), a newly recognised virus
disease of penaeid shrimp. J. Invertebr. Pathol, 42, 62-70.
7.
L I G H T N E R D.V.,
R E D M A N R.M.,
B E L L T.A.
& B R O C K J.A.
Detection of IHHN virus in Penaeus stylirostris and P. vannamei
into Hawaii. J. World Mariculture Society, 14, 212-225.
8.
L I G H T N E R D.V.,
REDMAN R.M.,
WILLIAMS R.R.,
(1983).
-
imported
M O H N E Y L.L.,
CLERX
J.P.M., B E L L T.Z. & BROCK J.A. (1985). - Recent advances in penaeid
virus disease investigations. J. World Mariculture Society, 16, 267-274.
9.
LIGHTNER D.V. & REDMAN R.M. (1989). - Hosts, geographic range and
diagnostic procedures for the penaeid virus diseases of concern to shrimp
culturists in the Americas. In: P. De Loach, W.J. Dougherty & M.A.
Davidson (eds). Frontiers of Shrimp Research. Elsevier, Amsterdam, pp.
173-196.
10.
L u Y., L O H P.C. & BROCK J.A. (1989). - Isolation, purification and
characterization of infectious hypodermal and hematopoeitic necrosis virus
(IHHNV) from penaeid shrimp../. Virol. Methods, 26, 339-344.
11.
M A R I J., BONAMI J.R. & LIGHTNER D.V. (1994). - Structure and cloning of
the genome of IHHNV, an unusual parvovirus pathogenic for penaeid
shrimp. Diagnosis of the disease using a highly specific probe. J. Gen.
Virol. (In press.)
12.
SEIDMAN C E . (1989). - Introduction of plasmid D N A into cells. In: F.M.
Ausubel, R. Brent, R.E. Kingston, D.D. Moore, J.G. Seidman, J.A. Smith
& K. Struhl (eds). Current Protocols in Molecular Biology, Vol. 1. John
Wiley and Sons, New York, pp. 1.8.1-1.8.8.
13.
VAGO C. & AMARGIER A. (1963). - Coloration histologique pour la
différenciation des corps d'inclusion polyhèdriques de virus d'insectes.
Ann. Epiphyties, 14(3), 269-274.
181
Yellowhead disease
C H A P T E R 25
YELLOWHEAD DISEASE
(No OIE number)
SUMMARY
Yellowhead virus (YHV), a non-occluded baculo-like virus, has been
demonstrated to be a potentially serious pathogen for the cultured
juveniles of Penaeus monodon. YHV is characterised by initiating
massive necrosis and vacuolated cells and hypertrophied nuclei of
infected tissues with densely stained basophilic inclusions
adjacent
to nuclei.
To date YHV epizootics have occurred only in Thailand, causing
very serious mass mortality. Experiments on artificial
infection
using the filtrate from the diseased shrimps showed a very high
mortality rate.
Two presumptive
tests, copper content in haemolymph
and
histopathological observations using light microscopy, may be used
for the diagnosis of YHV infection in shrimp. However,
definite
diagnosis should rely on transmission electron microscopy (TEM).
Under EM, enveloped virions with bacilliform morphology,
ranging
from 150-200 nm in length and 40-50 nm in diameter, have been
found in the cytoplasm of infected cells.
No eradication precedures have been developed for YHV infection.
Good pond management is so far the only prevention technology for
yellowhead disease (YHD). Some chemicals such as chlorine and
iodine have proved to be effective disinfectants. However, the use of
these chemicals
during the culture period of shrimp is not
recommended.
A. INTRODUCTION
Yellowhead disease (YHD) is a viral infectious disease of the giant tiger prawn,
Penaeus monodon. The causative agent has been demonstrated to be a nonoccluded baculo-like virus, yellowhead virus (YHV). It may initiate mass
mortality for cultured shrimps at the grow-out stage. To date, YHD has been
found in central and southern Thailand only.
Epizootics of yellowhead disease (YHD) of Penaeus monodon have occurred in
many areas of central and southern Thailand since 1990. The yellowhead virus
(YHV) infected moribund shrimps showed histopathological changes in gill
lamellae with extensive necrosis of pillar and epidermal cells and haemocytes.
Cellular necrosis, with hypertrophied nuclei and densely stained basophilic
182
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
globose cytoplasmic inclusions adjacent to nuclei, was also observed in
connective tissue underlying the midgut, in cardiac tissue, in haematopoietic
tissue, and in gill tissue. Results of the infectivity trials and field surveys
demonstrate that YHV may initiate a mass mortality for cultured P. monodon.
This virus seems to be the most virulent destructive virus so far detected in
cultured P. monodon.
B.
DIAGNOSTIC
TECHNIQUES
The moribund or heavily infected shrimps exhibit the clinical sign of a light
yellow colour in the céphalothorax and gills, that may easily be observed by the
naked eye. In addition, total haemocyte count, wet mount of gills, copper
content of haemolymph, Giemsa or haematoxylin staining of gills, and electron
microscopic observations are used for the diagnosis of YHD (3,4).
1.
W E T MOUNT TECHNIQUE
Infection of YHV infection can be rapidly diagnosed by the presence of
globular materials with a yellowish and fat-like structure in the secondary
lamellae of the gills.
2.
H I S T O P A T H O L O G I C A L OBSERVATION
Haemolymph collected from diseased shrimps is fixed in cold 10% formalin
in sea water. A thin film is then prepared on a glass slide and air dried.
Subsequently, Wright-Giemsa staining is performed on the blood film.
Under microscopy pyknotic and karyorrhectic nuclei of the haemocytes of
diseased shrimps are observed.
With light microscopy, sections of yellowhead specimens stained with
haematoxylin and eosin showed necrotic cells with hypertrophied nuclei and
large vacuoles. The disease was characterised by the presence of densely
basophilic, globose bodies adjacent to nuclei in the infected tissues in
lymphoid tissue, in the interstitial tissues of the hepatopancreas, in the
connective tissue underlying the midgut, in cardiac tissue, in haematopoietic
tissue and in gill tissue.
3.
C O P P E R CONTENT
HAEMOCYTES
OF
HAEMOLYMPH
AND
TOTAL
CELL
COUNT
OF
Haemolymph may be collected using a 1 ml plastic syringe and 20 G needle.
L-cysteine may be used as anticoagulant. The mixture of anticoagulant and
haemolymph (1:1) is immediately counted using a haemocytometer. The
results show that the haemocyte count of the YHV infected shrimps is
significantly lower than that of the normal ones. Haemolymph without
addition of anticoagulant is allowed to clot at room temperature and serum
is obtained following centrifugation at 6,500 rpm for 10 minutes. The
copper content of the serum may be analysed using atomic absorption
183
Yellowhead disease
spectroscopy. The results reveal that the copper content in haemolymph in
normal shrimps is significantly higher than that of the YHD infected
shrimps.
REFERENCES
1.
BOONYARATPALIN
S.,
SUPAMATTAYA
K.,
KASORNCHANDRA
I,
DlREKBUSARACOM S., A E K P A N I T H A N P O N G U. & CHANTANACHOOKLIN C.
(1993). - Non-occluded baculo-like virus, the causative agent of yellow­
head disease in the black tiger shrimp (Penaeus monodon). Gyobo Kenkyu
(Fish Pathology), 28, 103-109.
2.
CHANTANACHOOKIN
DlREKBUSRACOM
C,
S.,
BOONYARATPALIN
EKPANITHANPONG
S.,
KASORNCHANDRA
J.,
U.,
SUPAMATTAYA
K.,
SRIURAIRATANA S. & FIEGEL T . W . (1993). - Histology and ultrastructure
reveal a new granulosis-like virus in Penaeus monodon
yellowhead disease. Dis. Aquat. Org., 17, 145-157.
3.
affected by
EKPANITHANPONG U., BOONYARATPALIN S. & DIREKBUSRACOM S. (1993).
- Efficacy of calcium hypochlorite as a disinfectant against yellow-head
disease baculovirus in tiger shrimp (Penaeus
monodon).
Second
Symposium on Diseases in Asian Aquaculture, 25-29 October 1993, Karon
Villa Phuket Hotel, Phuket, Thailand.
4.
N A S H G . , ARKARAJAMORN A . & WITHAYACHUMNARNKUL B . (1992). -
Routine and rapid diagnosis of yellow-head disease in Penaeus
Asian Shrimp News, 4th Quarter,. 2-3.
5.
monodon.
SUPAMATTAYA K.J. & BOONYARATPALIN s S. (1993). - Comparative study
of simple methods for the diagnosis of yellow-head disease in the black
tiger shrimp (Penaeus monodon). Asian Shrimp News, 1st Quarter, Issue
No. 17, 2-3.
Crayfish plague
185
C H A P T E R 26
CRAYFISH PLAGUE
(No OLE number)
SUMMARY
Crayfish plague is a highly infectious disease of all crayfish
(Decapoda: Astacidae, Cambaridae) of non-North American origin.
The aetiological agent is an Oomycete fungus, Aphanomyces astaci,
which is now widespread in Europe as well as in North America. The
European crayfish species, the Noble crayfish Astacus astacus of
northwest Europe, the stone crayfish Austropotamobius pallipes of
southwest
and west Europe,
the related
Austropotamobius
torrentium (mountain streams of southwest Europe) and the slender
clawed or Turkish crayfish Astacus leptodactylus of eastern Europe
and Asia Minor are all highly susceptible. The only other crustacean
known to be capable of infection by A. astaci is the Chinese mitten
crab (Eriocheir sinensis) and this only under laboratory conditions.
The disease first occurred in Europe in the mid 19th century in
North Italy and then on the Franco-German border region. From the
latter region a steady spread of infection occurred, principally in
two directions - down the Danube into the Balkans and towards the
Black Sea, and across the North German plain into Russia and from
there south to the Black Sea and northwest to Finland and finally in
1907 to Sweden. In the 1960s the first outbreaks in Spain were
reported and in the 1980s further spread of infection to the British
Isles, Turkey, Greece and Norway have been reported.
The reservoir for the original infections in the mid 19th century was
never established, but the post-1960s extensions are largely linked
to movements of North American crayfish introduced more recently
for purposes of crayfish farming.
These species
(Pacifastacus
leniusculus [the Signal crayfish] and Procambarus clarki [the
Louisiana swamp crayfish]) can act as largely or
completely
asymptomatic carriers, but can be killed by A. astaci under adverse
conditions.
Transmission has also resulted from
contaminated
crayfish traps and other contaminated
equipment.
Clinically, infected crayfish may present a wide range of gross signs
of infection or none at all. Focal whitening of local areas of
musculature beneath transparent areas of thin cuticle, especially of
the ventral abdomen and in the periopod (limb) joints,
often
accompanied by even more localised brown melanisation, is the
most consistant sign. In the terminal stages of infection
animals
show a limited range of behavioural signs, principally a loss of the
normal aversion to bright light (they are seen in open water in
daylight) later accompanied by a loss of limb co-ordination
which
186
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
produces an effect which has been described as "walking on stilts".
Eventually, moribund animals lose their balance and fall onto their
backs before dying.
Diagnosis requires isolation and identification of the pathogen by
microscopic morphology; no biochemical or serological
methods
exist.
Control of spread of infection once a watershed is infected is in
practical
terms impossible. Prevention
of all introductions
of
crayfish to natural waters and into enclosed waters from which they
may escape to natural waters can be effective, although
movement
of fish can result in the movement of infected water between
watersheds
and can transmit infection
as can
contaminated
equipment such as boots and fishing gear. Malachite green has been
demonstrated to be effective in 'disinfecting' contaminated
water
and fish, and sodium hypochlorite and iodophores are effective for
disinfection
of contaminated
equipment.
Thorough
drying
of
equipment C>24 h) is also effective since the oomycetes are not
resistant to desiccation.
INTRODUCTION
Crayfish plague is a highly infectious fungal disease of fresh water crayfish. The
disease is of North American origin. North American crayfish species are
generally resistant to the disease, whilst all crayfish species from other
continents are highly susceptible. The causative agent is an Oomycete fungus,
Aphanomyces astaci. Other members of this genus are now implicated in a
number of fish diseases, particularly the epizootic ulcerative syndrome of South
East Asia.
The disease first appeared in Europe in about 1860. By 1935 only the British
Isles, the Iberian Peninsula, Greece, Turkey and Norway remained free of
infection. In the period 1960-1980 those areas were also infected. In most cases
evidence exists to link these more recent extensions to the transfer of infected
carrier crayfish introduced for farming.
The first sign of a crayfish plague mortality may be the presence of numbers of
crayfish at large during daylight (crayfish are normally nocturnal), some of
which show evident loss of coordination in their movements and easily fall over
onto their backs and are unable to right themselves. Often, however, unless
waters are carefully observed, the first recognition that there is a problem will
be the presence of large numbers of dead crayfish in a river or lake.
In susceptible species where sufficient numbers of crayfish are present to allow
infection to spread rapidly, and particularly at summer water temperatures,
infection will spread quickly and stretches of over 50 km of river may lose all
their crayfish in less than 21 days from the first observed mortality. Crayfish
Crayfish plague
187
plague has an unparallelled severity of effect; infected susceptible crayfish do
not survive - 100% mortality is the norm. Resisitant North American species do
survive infection in many cases and then act as largely asymptomatic carriers,
although under adverse conditions (stress, other infections), a major crayfish
plague mortality may occur.
Infection is transmitted horizontally through river water by means of the fungus'
motile biflagellate zoospores, which exhibit a positive chemotaxis towards
crayfish. Depending on the site of the initial source of infection, the mortality
will spread downstream at the speed of flow of the river, and upstream at 2 to 4
km per year. The upstream spread may be attributed to normal movements of
crayfish between infection and the development of clinical disease.
Infection may also be transmitted in transport water (e.g. with movements of
fish between farms) and on contaminated equipment (boots, fishing gear, etc.).
Fish and fish water may be disinfected by standard therapeutic doses of
malachite green during transport, and equipment with hypochlorite or
iodophores. No therapy of infected crayfish is possible.
Diagnosis of crayfish plague strictly requires the isolation and characterisation
of the pathogen, A. astaci, using simple mycological media fortified with
antibiotics to control bacterial contamination. Isolation is only likely to be
successful before or within 12 hours of the death of infected crayfish. However,
there is no other disease or pollution effect which can cause such total
mortalities of crayfish, while leaving all other animals in the same water
unharmed, so that isolation of the pathogen is desirable but not essential,
particularly in regions where further spread of infection is known to be a
potential hazard. Clinical signs of crayfish plague include behavioural changes
and a range of visible external lesions. The range of these is however very large
so that, except for the experienced eye, such clincal signs are of limited
diagnostic value.
B.
DIAGNOSTIC
I S O L A T I O N AND I D E N T I F I C A T I O N O F ,4.
PROCEDURES
ASTACI
Isolation methods are as described by Alderman and Polglase. An agar medium
(IM) is used, containing yeast extract and glucose in river water with
antimicrobial agents (penicillin G and oxolinic acid) which prevent the growth
of most bacteria and enable easy and rapid isolation of the pathogen.
I M medium:
Agar
Yeast extract
Glucose
Oxolinic acid
River water
Penicillin G (sterile) added after
autoclaving and cooling to 40°C
12.0 g
1.0 g
5.0 g
10 mg
1000 ml
1.0 g
188
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Simple aseptic excision of infected tissues which are then placed as small
pieces (1-2 m m ) on the surface of IM plates will normally result in
successful isolation of A. astaci from moribund or recently dead (<24 h)
animals. On IM agar growth of new isolates of A. astaci is almost entirely
within the agar except at temperatures below 7°C, when some superficial
growth occurs. Colonies are colourless. Dimensions and appearance of
hyphae are much the same in crayfish tissue and in agar culture. Vegetative
hyphae are aseptate (5)7-9(10) um in width (i.e. normal range 7-9 um, but
observations have ranged between 5 and 10 um). Young, actively growing
hyphae are densely packed with coarsely granular cytoplasm with numerous
highly refractile globules. Older hyphae are largely vacuolate with the
cytoplasm largely restricted to the periphery with only thin strands of
protoplasm bridging the large central vacuole. The oldest hyphae are
apparently devoid of contents. Hyphae branch profusely, with vegetative
branches often tending to be somewhat narrower than the main hyphae for
the first 20-30 um of growth.
2
When actively growing thalli or portions of thalli from broth or agar culture
are transferred to distilled water, sporangia form readily in 20 to 30 h at
16°C and 12 to 15 h at 20°C. Sporangia are myceloid, terminal or
intercalary, developing from undifferentiated vegetative hyphae. Sporangial
form is variable: terminal sporangia are simple, developing from new
extramatrical hyphae, whilst intercalary sporangia can be quite complex in
form. Intercalary sporangia develop by the growth of a new lateral
extramatrical branch which forms the discharge tube of the sporangium. The
cytoplasm of such developing discharge tubes is noticeably dense and these
branches are slightly wider (10-12 um) than ordinary vegetative hyphae.
Sporangia are delimited by a single basal septum in the case of terminal
sporangia and by septa at either end of the sporangial segment in intercalary
sporangia. Such septa are markedly thicker than the hyphal wall and have a
high refractive index. Successive sections of vegetative hypha may develop
into sporangia and most of the vegetative thallus is capable of developing
into sporangia.
Within developing sporangia the cytoplasm cleaves into a series of elongate
units (10-25 x 8 um) which are initially linked by strands of protoplasm.
Although the ends of these cytoplasmic units become rounded, they remain
elongate until and during discharge. Spore discharge is achlyoid, that is, the
first spore stage is an aplanospore which encysts at the sporangial orifice
and probably represents the suppressed saprolegniaceous primary zoospore.
No evidence has been observed for the existence of a flagellated primary
spore, thus, in this description, the terms 'sporangium' not 'zoosporangium'
and 'primary spore' not 'primary zoospore' have been used. Discharge is
fairly rapid (<5 min) and the individual primary spores (=cytoplasmic units)
pass through the tip of the sporangium and accumulate around the
sporangial orifice. Speed of cytoplasmic cleavage and discharge is
temperature dependent. At release, each primary spore retains its elongate
irregularly amoeboid shape briefly before encystment occurs.
Crayfish plague
189
Encystment is marked by a gradual rounding up followed by the
development of a cyst wall which is evidenced by a change in the refractive
index of the cell. From release to encystment occupies 2 to 5 min. Some
spores may drift away from the spore mass at the sporangial tip and encyst
separately. Formation of the primary cyst wall is rapid and once encystment
has taken place the group of spores remains together as a coherent group
and adheres well to the sporangial tip so that marked physical disturbance is
required to break up the spore mass.
Encysted primary spores are spherical (8)9-11(15) pm in diameter and are
relatively few, (8)15-30(40) pm per sporangium in comparison to other
Aphanomyces
spp. Spores remain encysted for 8-12 h. Optimum
temperatures for sporangial formation and discharge lie between 16 and
24°C, but the discharge of secondary zoospores from the primary cysts peaks
at 20°C and does not occur at 24°C. In new isolates of A. astaci, it is normal
for the majority of primary spore cysts to discharge as secondary zoospores,
although this varies with staling in long term laboratory culture. Sporangial
formation and discharge occurs down to 4°C.
In many cases, some of the primary spores are not discharged from the
sporangium and many sporangia do not discharge at all. Instead, the
primary spores appear to encyst in situ within the sporangium, often develop
a spherical rather than elongate form and certainly undergo the same
changes in refractive index that mark the encystment of spores outside the
sporangium. Such within sporangial encystment has been observed on
crayfish. Spores encysted in this situation appear to be capable of
germinating to produce further hyphal growth.
Release of secondary zoospores is papillate, the papilla developing shortly
before discharge. The spore cytoplasm emerges slowly in an amoeboid
fashion through a narrow pore at the tip of a papilla, rounds up and begins a
gentle rocking motion as flagellar extrusion begins and spore shape changes
gradually from spherical to reniform. Flagellar attachment is lateral;
zoospores are typical saprolegniaceous secondary zoospores measuring 8x12
pm. Active motility takes some 5-20 min to develop (dependent on
temperature) and, at first, zoospores are slow and uncoordinated. At
temperatures between 16 and 20°C, zoospores may continue to swim for at
least 48 h.
REFERENCES
1.
2.
ALDERMAN D . J . & POLGLASE J . L . (1986). - Aphanomyces
and culture. J. Fish Dis., 9, 367-379.
astaci: isolation
ALDERMAN D . J . , POLGLASE J . L . & FRAYLING M . (1987). -
Aphanomyces
astaci pathogenicity under laboratory and field conditions. J. Fish Dis., 10,
385-393.
Lists
191
1. LIST OF OIE REFERENCE LABORATORIES
FOR FISH, MOLLUSC AND CRUSTACEAN
DISEASES
2. LIST OF ORGANISATIONS WITH WHICH THE
OIE HAS COOPERATION AGREEMENTS
193
Lists
1. L I S T O F OLE R E F E R E N C E L A B O R A T O R I E S F O R F I S H ,
M O L L U S C A N D C R U S T A C E A N D I S E A S E S LN 1 9 9 5 *
Diseases/Viruses
Viral haemorrhagic
septicaemia virus
Virus de la septicémie
hémorragique virale
Expert/Laboratory
Dr N. J ô r g e n Olesen
Statens Veterinaere Serum Laboratorium, 2 Hangovej, DK8200 Aarhus N, DENMARK
Tel: (45) 86.16.79.00, Fax: (45) 86.10.74.64
Virai de la septicaemia
hemorrágica viral
Spring viraemia of carp
virus
Virus de la virémie
printanière de la carpe
Dr B.J. Hill
Fish Disease Laboratory, MAFF, Barrack Road, The Nothe,
Weymouth, Dorset DT4 8UB, UNITED KINGDOM
Tel: (44) 1305.20.66.00, Fax: (44) 1305.20.66.01
Virus de la viremia
primaveral de la carpa
Infectious haematopoietic
necrosis virus
(Rhabdoviruses)
Virus de la nécrose
hématopoïétique
infectieuse
(Rhabdoviroses)
Virus de la necrosis
hematopoyética infecciosa
(Rhabdoviruses)
Oncorhynchus
virus
masou
Virus de 1' Oncorhynchus
masou
Virus del
masou
Dr Jo-Ann L e o n g
Oregon State University, Department of Microbiology, Nash
Hall 220, Corvallis, Oregon 93331-3804, USA
Tel: (1.503) 737.4441, Fax (1.503) 737.0496
Dr J . W i n t o n
Fish and Wildlife Service, National Fisheries and Research
Center, Building 204, Naval Station, Seattle, Washington
98115-5007, USA
Tel: (1.206) 526.6587, Fax: (1.206) 526.6654
Dr M. Y o s h i m i z u
Hokkaido University, Faculty of Fisheries, Hakodate, Hokkaido
014, JAPAN
Tel: (81.138)41.0131, Fax: (81.138)43.5015
Oncorhynchus
Epizootic haematopoietic
necrosis virus
Virus de la nécrose
hématopoïétique
épizootique
Dr A. Hyatt
Australian Fish Health Reference Laboratory, c/o Australian
Animal Health Laboratory, P.O. Bag 24, Geelong, Vic. 3220,
AUSTRALIA
Tel: (61)52.27.5000, Fax: (61)52.27.5555
Virus de la necrosis
hematopoyética epizoótica
Continued overleaf
*
This list is updated annually and the revised list will be published in the
May issue of the OIE Bulletin.
194
OIE Diagnostic Manual for Aquatic Animal Diseases, 1995
Reference Laboratories, contd.
Mollusc pathogens
Agents pathogènes des
mollusques
Dr H. Grizel
IFREMER, Laboratoire de Pathologie des Invertébrés, 17390
La Tremblade, FRANCE
Tel: (33) 46.36.30.07, Fax: (33) 46.36.37.51
Agentes patógenos de los
moluscos
Crustacean pathogens
Agents pathogènes des
crustacés
Agentes patógenos de los
crustáceos
Dr D. Lightner
Department of Veterinary Science, Building 90, Room 202,
Tucson AZ 85721, USA
Tel: (1.602) 621.6903, Fax: (1.602) 621.6366
Prof. S.N. Chen
Department of Zoology, Director, Institute of Fishery Biology,
National Taiwan, University, No. 1 , Roosevelt Road, Section 4,
Taipei, Taiwan
TAIPEI CHINA 10764
Tel: (886.2) 368.71.01, Fax: (886.2) 368.71.22
195
Lists
2 . LIST OF ORGANISATIONS WITH W H I C H THE OIE
HAS COOPERATION AGREEMENTS
The OLE has signed a number of agreements with international organisations
having an interest in animal health or veterinary public health. These
agreements aim to ensure that there is a coordination of effort and cooperation
in carrying out joint actions where appropriate. The organisations concerned are
the following (when two dates are given, these refer to the different dates on
which the agreement was officially ratified by the two organisations):
1.
Food and Agriculture Organisation of the United Nations (FAO)
Agreement signed in 1952/1953
2.
W o r l d Health Organisation ( W H O )
Agreement signed in 1961/1962
3.
Inter-American Institute for Cooperation on Agriculture
Agreement signed in 1981
4.
P a n A m e r i c a n Health
(PAHO)
Agreement signed in 1993
Organisation/World
Health
(nCA)
Organisation