and Biopsy Procedures
Written by Robert B. Moeller Jr., DVM
Fish, like other vertebrates, have a complex nervous system and when
handled, can experience stress. When performing most diagnostic
procedures, fish should be anesthetized before handling. If a fish is to
be submitted for pathology, euthanasia should be done prior to placing the
animal into 10% formalin or other fixative. It would be considered
inhumane to place a fish in 10% neural buffered formalin or other fixative
The 1993 AVMA Report of the AVMA Panel on Euthanasia states that
acceptable anesthetics to be used for the euthanasia of fish are tricane
methane sulfate, benzocaine and barbiturates. A conditionally acceptable
method for euthanasia is a blow to the head followed by decapitation.
Other commonly used methods are carbon dioxide (four alka-selzer tablets
to 500 mls of water), electrocution and hypothermia. Other anesthetic
agents are available for immobilizing fish for euthanasia; an excellent
review of these agents and their mechanism of action can be found in
Stoskopf's book, Fish Medicine (1993).
The preferred method of euthanasia would be to anesthetize the fish to
a deep plane of anesthesia (stage III or stage IV) and then sever the
spinal cord just caudal to the brain. The most common and practical way to
anesthetize a fish is to place the anesthetic agent in water. The fish
will go through all four stages of anesthesia prior to death. The four
stages of anesthesia and the clinical presentation of each are as follows:
Stage I; Induction and light
sedation: The fish goes through an excitement phase with erratic swimming
followed by reduced activity. The respiratory rate increases and there is
a loss of some response to tactile stimulation.
STAGE II; Sedation: Fish swim
slowly, have decreased gill movement (respiration), and a loss of
STAGE III; Anesthesia: Fish have a
complete loss of equilibrium and are unable to swim. Gill movement
(respiration) becomes very slow. The fish is unresponsive to external
Stage IV; Anesthetic overdose: The
fish has a total loss of gill movement and the opercules become distended.
The fish goes into cardiac arrest.
Collecting Samples for Bacteria and Fungi
Sampling for bacteria and fungi should be done on fish that are brought
in for examination alive or from fresh fish that had died only recently
(usually less than 6 hours).
Ideally, the fish is alive when sampling cutaneous lesions. The area to
be cultured should not be handled prior to culturing. Like cutaneous
smears, large ulcerated areas should be avoided and small developing
lesions cultured. The sterile loop or culturette should be rubbed into the
As with cutaneous lesions, gills from live fish are the best to
culture. Gills are cultured by gently rubbing and rolling the sterile
culturette through the gill arches. The culturette should pick up abundant
mucus on the tip of the culturette. Since the gills and cutaneous lesions
are exposed to the aqueous environment, a mixed bacterial culture should
be expected. If a pure culture of a potential pathologic bacteria is
obtained, this should be considered a possible cause of disease in this
The kidney is one of the most important internal organs to culture.
There are two methods of culturing the kidney. The first method is to cut
the dorsal fin off, sterilize the open area with heat, cut the vertebra
with a sterile scissors or scalpel and snap the fish by bringing the head
and tail together. This exposes the kidney for culturing. The second
method, can allow for possible contamination of internal organs. Here the
fish is dipped into 70% alcohol and the abdominal cavity opened
aseptically to allow all organs to be exposed. Sterilize with heat, the
desired internal organs to be cultured, cut open the organ with a sterile
scalpel and culture.
Another method for culturing, particularly in cases where a bacterial
septicemia is suspected, is to examine heart blood. Collection of heart
blood from the atrium is most productive since the atrium contains
phagocytic cells that assist in clearing bacteria from the blood.
For biopsy specimens and bacterial cultures, a fish does not need to be
killed. The fish should be anesthetized prior to performing most clinical
examinations and biopsies. Biopsy procedures on fish usually are cutaneous
smears, fin biopsies and gill biopsies.
Cutaneous smears are done primarily for ectoparasites. Large ulcerated
lesions should be avoided; Try to find smaller developing lesions for
sampling. Prior to performing cutaneous smears, bacterial cultures should
be taken. The procedure for cutaneous smears involves passing several
clean microscope slides over the area of interest. Only light pressure on
the glass slide needs to be used to remove some epidermis and mucus. On
one slide, a drop of water is placed on the smear and a cover slip is
placed on the slide for examination. The other slides should be air dried
or fixed in alcohol. These are stained with either new methyl blue stain
or Diff-Quick stain.
A fin biopsy is accomplished by spreading out the fin and a triangular
wedge shaped piece of tissue is cut between the rays of the fin. Place the
fin biopsy on a slide with water and cover slip for examination.
A gill biopsy is performed by cutting a few tips of the primary lamella
with the blades of the scissors. Place the lamellar tips on the slide with
a drop of water, cover slip and examine. Both fin and gill biopsies should
not cause undue harm to a fish.
Ideally, the fish should be submitted alive for the post mortem
examination. This gives the pathologist a chance to observe the fish prior
to euthanasia and note any important clinical signs. Unfortunately, some
situations do not allow the pathologist to evaluate the fish while they
are alive. Fish should be dead less than 6 hours. Fish found floating in a
tank longer than 6 hours are poor candidates for necropsy due to post
mortem autolysis. Dead fish should be wrapped in paper or gauze and
refrigerated. Do not freeze the fish.
Prior to performing the necropsy, insure that all necropsy tools,
sterile loop or culturettes, glass slides for impression smears, 70%
alcohol and 10% neutral buffered formalin or Bouin's solution (I prefer
Formalin) are available. A systematic approach should be used when
performing the necropsy. Evaluate the external surface and note the
general body condition of the fish, identify and note lesions on the skin,
fins, eyes, oral cavity and anus. Take cultures of the desired lesions.
After completion of the external examination, place the fish in lateral
recumbency on a disposable towel. Remove the eyes and then the operculum
with the pseudobranch, place these in formalin. Remove the second and
third gill arches being careful not to crush the primary lamella. Take
several primary lamella from one of the gill arches and place on the glass
slide for parasitic examination. Place the remaining gills into the
Using aseptic techniques, open the abdominal cavity by cutting through
the pectoral girdle to the spine and follow the abdominal cavity to the
anus, extend this cut along the ventral midline from the gills to the anus
and remove the body wall. Remove the body organs (heart, liver,
intestines, spleen, gonads and swim bladder) for examination. When
submitting the swim bladder for histopathology, insure that the red gas
forming organ is present. Sample both the anterior and posterior kidney in
fish with both kidneys. In fish with fused kidneys, insure that anterior
and posterior sections are submitted for examination.
Remove the brain by opening the skull just dorsal to the eyes and
removing the bones over the brain. Sample all cutaneous lesions for
histopathology. Insure that normal tissue from the margin of the lesion
are submitted with the lesions. Cut into the skeletal muscle and look for
parasitic cysts. Finally, open the stomach and intestine and examine food
If toxicology is desired, be sure to submit gills, kidney, liver,
skeletal muscle, and fat. Toxicologic samples should be immediately frozen
and stored in at -70 degrees C. Analysis of the tissue should occur as
soon as possible after collection.
DISEASES OF FISH
Robert B. Moeller Jr., DVM
California Animal Health and Food Safety Laboratory System
University of California
18830 Road 112
Tulare, California 93274
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