Diagnosis of Internal organs of fish. Anatomy of fish. Step by step procedure to handle and dissect the internal organs and sample collection for disease diagnosis.

1. Dissection of
Internal Organs of Fish
Submitted by Submitted to
B.Naveen Rajeshwar Dr TJ Abraham
Dept of AAH Dept of AAH

2. Internal Anatomy of Fish

3. Collecting fish
 Note: Taking fish or fish samples in excess of the minimum required is
recommended because further investigations may not be possible if
insufficient samples are taken.
 If dissection is not possible within 24 hours, whole fish may be frozen.
 Determine the number of fish to be collected, and which tissue is
required from each fish for the analyses to be conducted.
 A sample should have at least 60 fish of the same species of
approximately uniform size per site if possible.

4.  Collecting appropriate samples:
 In order to obtain fresh tissue after a fish kill, it is preferable to choose fish
that are sick or dying rather than dead (e.g. some might be moving but
showing signs of lethargy or distress).
 When sampling live fish, ensure that the fish are handled and euthanized
humanely (such as with the use of MS222).
 If only dead fish are present, choose the least decomposed fish available.
 Place individual samples into individual resealable bags with a label stating
relevant information such as date, time, sampler details, site, species and
replicate number.
 Place samples with crushed ice to transport to a laboratory, or clean area if
dissections are to be carried out in the field.

5. Preparing for dissection
► Ensure there is a clean working area and equipment can be rinsed between each
sample.
► Clean sampling equipment to be used for the dissection:
► Tools, work surface, and sample containers must be clean and not likely to
contaminate the samples of interest
► After each fish is dissected, all equipment should be cleaned and rinsed, and the
cutting board covering and gloves need to be changed.
► Clear an area to conduct the dissection.

6. ► Set up the work area to ensure all equipment is easily accessible once
dissections begin.
► Place a waste bin in an area easily accessible to the person conducting the
dissections.
► Identify a procedure for naming each sample/replicate/organ, and relating these
back to the individual sample.

7. Fish dissection
 Measure and weigh fish in accordance with the Fish holding,
identification and measurement of length and weight document. Record
details.
 Put on powder-free gloves. Gloves must be stored in a clean environment
(e.g. in a resealable plastic bag).
 Lay fish flat on one side with the dorsal fin facing away from you.

8. Gill samples
If the gills are to be collected:
 Lift the operculum (gill cover) and cut this off at its base to expose the
gills. Take care not to damage the gills when doing this.
 Carefully cut out the gills at their base Rinse gills with de-ionised water.
 Place gills in labelled storage container/bag
 Note: Gills on larger fish may not require the operculum to be removed.

9. Lifting the operculum (gill cover) Cut at the base

10. Muscle samples
 Muscle (flesh) samples should be collected above the lateral line, between the
dorsal fin and the caudal fin.
 This will maximise the amount of muscle tissue collected and reduce the risk of
accidentally piercing internal organs.
 Avoid cutting below the fish’s lateral line to ensure the lower intestine or other
internal organs are not pierced.
 If the intestine is cut open, this will lead to contamination of the organs and the
sample will not be usable.
 If muscle is to be collected, Make a cut with the scalpel blade from just below
the start of the dorsal fin down to the fish’s lateral line

11. Outline of area to be removed from the fish for
muscle sample
Removing skin
Removing muscle

12.  Cut from just above the lateral line of the fish toward the tail.
 Cut from where the first incision was made just below the dorsal fin across the
top of the fish and down toward the tail to meet the cut from step 2.
 Remove the skin of this section of cut flesh using forceps and a scalpel blade
Take care not to touch this exposed muscle.
 To remove the muscle sample, make incisions around the dissected area, cutting
underneath the flesh to detach it from the small bones and allow it to be
removed.
 Once the muscle has been removed from the fish, rinse it in deionised water.
 Place muscle sample in labelled storage container/bag

13. Internal organ samples
 1)Incision at anus- Begin by inserting a fine scalpel blade into the anus (also
called the vent) of the fish. The anus is located just anterior to (in front of) the
anal fin, on the ventral (lower) side of the fish in most fishes.
 2. Cutting anteriorly. The incision is then extended anteriorly along the fish's
belly towards the head.
 3. Cut between pelvic fins. The incision passes anteriorly between the pelvic
(ventral) fins. Depending on the type of fish, these paired fins are used to
stabilise the fish when swimming and also for braking. The pelvic fins are
supported by the bones of the pelvic girdle which are anchored in the belly
muscles.

14. Step: 4
Step: 2
Step: 1
Step: 3

15.  4. Cut along isthmus. Use scissors to cut anteriorly through the bones attached to
the pelvic fins. Cut forward along the narrow, fleshy space beneath the head and
between the gill covers. The gill covers (also known as operculae) are flaps which
lie along both sides of the head and protect the underlying gills.
 5. Body cavity. Pull apart the two walls of the body cavity and expose the internal
organs (see next image for names). The neat incision now runs from the anus
forward between the two pelvic fins and along the isthmus.
 6. Internal organs. Some of the ventrally located internal organs: 1 heart, 2 Liver,
3 Pyloric caecae, 4 adipose (fatty) tissue.
 7. Pull aside gut. Here the adipose tissue (1) and gut (2) are pulled aside to expose
the swim bladder (3), gonads (4) and kidneys (5). As a general rule, carnivorous
fishes have short guts. Herbivorous fishes have much longer guts. The gonads and
kidneys are paired. One of each can be seen on both sides of the swim bladder.

16. Step: 7
Step: 5
Step: 8
Step: 6

17.  8. Cut posterior end of gut. The gut is severed at the posterior end of the body
cavity, near the anus. The gut and other organs attached to it are pulled forward out
of the way, or removed entirely.
 9. Pull gut forward. Pulling the gut forward exposes the swim bladder (1), gonads
(2) and kidneys (3) in position dorsally (at the top) in the body cavity. A larger
portion of the liver is now visible (4). The kidneys are paired organs located in the
body cavity ventral to (below) the vertebral column. They are one of the organs
involved in excretion and regulation of the water balance within the fish.
 10. Swim bladder exposed. The other organs have been removed to expose the
swim bladder at the top of the body cavity. The swim bladder (also called the gas
bladder or air bladder) is a flexible-walled, gas-filled sac located in the dorsal
portion of body cavity. This organ controls the fish's buoyancy and is used for
hearing in some species.

18. Step: 9 Step: 10

19.  Note: Organs can be located in differing/varying places depending
upon the body shape of the species The kidney is a relatively
difficult organ to locate and dissect successfully. It is usually located
up close to the spine and may be hidden by the swim bladder.
 Rinse the removed organ with de-ionised water.
 Place the removed organ in labelled storage container/bag.

22. Preserving and packing samples
 Packing samples will depend upon the analysis required and
should be discussed with the laboratory prior to dissections.
Individual organs should be separately packaged, labelled
and preserved prior to sending to the laboratory.

23. Bacterial Infections
 Collection of samples for bacteriological examination should be taken
immediately after the fish is opened to minimize contamination. Samples for
detecting bacterial infections are routinely taken from the kidney. If lesions are
apparent in other tissues, sample those tissues using the same procedures.
 First expose the kidney by pulling the intestine away from the swim bladder,
and then gently strip the swim bladder from the kidney surface.
 Inoculate a sample of kidney onto bacteriological media as follows:
1. Flame a metal loop until red hot, and then insert it into the anterior part of the
kidney. The sample is immediately inoculated onto a plate of sterile agar by
streaking back and forth across the surface of the agar at one end of the plate.

24. 2. Streak for isolation by reflaming the loop, then allow it to cool and make several streaks
into a new area of the plate. This is repeated three to four times to insure the bacteria is
diluted over the surface of the agar.
3.Label the bottom of the plate and incubate with the agar surface on top (i.e., upside
down) either in an incubator or on a bench top.
 Bacterial colonies will normally be visible within 48 to 72 hours of inoculation. The
shape and pigmentation of the colonies should be noted. Varied types of colonies, and
colonies that grow outside of the streak lines, usually indicate that your bacterial
sample has been contaminated.

25. References
 Fish collection and dissection for the purpose of chemical analysis of tissues,
Biological assessment, 2018.
 Fish dissection guide
Thank You…….!