Appendix 2: Post-mortem Procedures

Appendix 2: Instructions to pathologists

FINFISH

1. Purpose and Scope

The purpose of this procedure is to provide a guide to laboratory staff conducting finfish post mortems to enable a consistent approach. This procedure applies to all finfish post mortems.

2. Procedure

The following post mortem procedure describes a basic technique for finfish.

External examination

• The surface of the necropsy table, dissecting board and equipment are cleaned and treated with Virkon^® or ethanol between cases or animals to reduce contamination of the working area.
  
• Pots containing buffered formalin 10% (diluted in seawater or freshwater with buffering salts) sterile bacteriology sample containers/bottles, syringes/needles and swabs expected to be used during the post mortem are labelled with the Case number prior to commencing.
  
• All equipment needed to perform the post mortem is placed together including forceps, scalpel, scissors and any other equipment necessary. The size of the sample will dictate other necessary equipment.
  
• The wearing of a plastic apron, enclosed shoes and gloves is mandatory for all staff carrying out post mortems.
  
• After euthanasia, place the fish on the dissecting board or in a shallow dish where necessary. Convention states that fish are drawn/photographed from the left lateral view (i.e. with the head to the left).
  
• A worksheet is used to record each batch of specimens and for each individual specimen when greater detail is required.
  
• Systematically check the outer surface, fins, eyes, nares (nostrils), oral cavity and under the operculum. Look at the gills in situ and note their colour and any discoloured areas or ragged appearance. Spread the fins and check for parasites.
  
• Skin: Scrape the skin in the direction of the scales (craniocaudal). This is best done behind the pectoral fin and on any visible lesions and examine under low power with a drop of water. Motile protistan parasites and monogenean flukes are common. Giemsa or Gram stains may be performed. NOTE: Skin scrapings can be done on live fish.
  
• Gills: Lift the operculum or remove with scissors. Larger gill samples may be removed and examined under low power in a Petri dish of water. Examine a few primary lamellae under high power. Motile protistan parasites and monogenean flukes are common.

NOTE: A gill biopsy can be performed on a live fish over 3 cm in length. The operculum is lifted with forceps and the tips of 2-3 gill filaments snipped off with fine scissors. Haemostatis is instantaneous. Examine gill specimen in a drop of water.

• Blood: Sample may be taken from the caudal vein or the sinus (ductus Cuvieri) behind the posterior wall of the opercular cavity, or from the heart through the body wall.
  
• Eyes: Examine for parasites and exophthalmos. Eyes may be removed to check for haemorrhage, gas or oedema fluid or other abnormality behind the eyeball.
  
• Nares: Cut nares (nostrils) and examine a smear of the lining.
  
• Anus: Check for distension, reddening, ulceration or discharge.

Internal examination

• Make a cut along the ventral surface immediately anterior to the anus. Insert scissors and cut forward to the base of the pectoral fins. Cut through the pectoral girdle and then cut up the edge of the operculum to the top of the abdominal cavity and then back towards the vent (ensuring that the internal organs are not ruptured) so that the flap of the body can be removed. Refer to Figure 1

^{Figure 1: Representation of the incision line for internal examination.}

• Check for ascites. Observe grossly the liver, spleen, heart, stomach, intestines, abdominal fat, swim bladder, peritoneum, muscles and kidney. Check abdominal organs for colour, friability, adhesions, haemorrhages, necrosis, nodules, cysts and parasites. Incisions should be made in muscle blocks to check for haemorrhage and parasites. Refer to Figure 2.

^{Figure 2: Internal organs of a bony fish.}

• To check for endoparasites, remove the internal organs into a Petri dish, open the gastrointestinal tract along its length, check the presence or absence of food and any lesions on the walls. Scrapings should be examined for metazoan and protistan parasites by wet smear. Intestinal contents can be sieved; the size necessary will depend on the specimen. Observe the size and colour of the gall bladder. Smear the contents of the gall bladder.
  
• To examine the brain, cut into the cranium, posterior to the eyes with a bone-cutting instrument. The brain in finfish is small and requires careful extraction. When the cavity is exposed, lift the brain out with forceps (curved forceps seem most useful) by placing them under the entire brain and pulling in an upwards motion.

Sample collection

Normal diagnostic:

Collection of kidney, liver, spleen, thymus, gill, muscle, brain, blood, lesions and intestine.

Specialised sampling

• Bacteriology: Take the relevant samples.
  
• Electron Microscopy: Take the indicated samples.

Sampling tissues from a finfish post mortem - for histology

Introduction

The essential requirements for sampling are:

• That the specimen must be placed in the chemical fixative as quickly as practicable (especially the gill tissue, as this begins to alter several minutes post death) and ensure that the tissues have not been previously frozen.

Note the elapsed time of death, if known, and the state of the sample on the work sheet.

• Specimens are in sufficient fixative (10:1 v/v or greater).
  
• Specimens are handled gently to prevent artefacts and damaging architecture.
  
• That the tissues are trimmed to dimensions of 10 mm by 3 mm thick. Small fish can be fixed whole, provided a longitudinal slit is made in the abdominal cavity to expose internal organs to the fixative.

Lesions

When sampling gross lesions select a piece of tissue that includes normal and affected zones.

Decalcification

Tissue samples containing calcium in the form of bone or scales should be fixed, trimmed and then submitted to the Histology section with a note to decalcify prior to processing.

Skin with scales- cut oversize, decalcify and then trim to size for histology (scales are easily dislodged by cutting in fresh material)

Fixatives

The fixatives recommended are 10% neutral buffered formalin or 10% formalin in seawater.

Sampling tissues from a finfish post mortem for Bacteriology

Introduction

The three essential requirements for sampling are:

• Blood and swab samples should be taken aseptically.
  
• That bacteriological isolation is dependent on the correct choice of growth media. This choice is determined by the environment from which the specimen was taken (temperature, salinity etc.).

Samples submitted for bacterial culture

Blood sampling

The removal of blood from circulation can be done in several ways: truncation of the caudal peduncle (tail), heart puncture, caudal vessel puncture, ductus Cuvieri puncture and repeated sampling via chronic cannulation of the aorta. Refer to Figure 1.

^{Figure 1. Common sites for obtaining blood samples from fish: A, heart; B, puncture of the caudal vessels; C, puncture of the Ductus Cuvier.}

Destructive sampling

Blood may be collected via any method after ensuring that the area of puncture is flooded with 70% ethanol. The most favoured site are the caudal vessels or the ductus Cuvieri (Figure 1). Caudal truncation is usually only performed on small fish since the finer anatomical features render other techniques less efficient. Cardiac puncture is most easily performed by exposing the heart so that needle placement can be precise.

Non–Destructive sampling

If a blood sample only is required, the sacrifice of the fish is not necessary. The use of effective non–lethal doses of anaesthetics and a puncture technique allow the sampled fish to be returned to the population upon recovery.

The choice of sampling site is influenced by several considerations: the thickness of the overlying tissue, the ease of exact location of the site from anatomical landmarks and the potential for damage to the vessels and associated tissues.

The site of choice is the puncture of the caudal vessels. See Figure 1.

Liver

A slice of liver is submitted at least 3 mm thick, enabling the sample to be flooded with ethanol and flamed before a swab is plunged in the centre. Alternatively, place the sample into a Petri dish.

Spleen

A slice of spleen can be sampled in the same way as the liver described above.

Lesion

Lesions can be cut from the animal and treated the same as other tissues, taking a swab sample or in the case of a skin lesion. With small fish the whole animal can be flooded with ethanol, flamed and a swab plunged into the lesion.

Kidney

A slice of kidney can be removed, and treated in the same way as the liver sample mentioned above. The preferred method for sampling is to pierce the swim bladder and flood with ethanol. Remove the excess ethanol by rolling the liquid off the site of puncture. Puncture the kidney on a dorso-ventral angle, drawing both blood and tissue. The anatomy of the kidney will depend on the species of fish being sampled. The site of puncture should be chosen to have the largest tissue mass. Smaller fish can be sampled by cutting anterior to the caudal peduncle, (ensuring that the abdomen stays intact) flooding the cut surface with ethanol and placing the needle directly into the kidney.

Samples submitted for fungal culture

If swabs are to be taken for fungal culture of infected sites, then this must be done prior to any of the abovementioned aseptic techniques.

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