Necropsy

The opercula may be injured, shortened, deformed or eroded (see FPA). Raise the operculum using a forceps (“rat tooth” forceps work well), to observe the cavity and branchial arches. You may need to cut the operculum vertically using pincers or scissors, as needed, to facilitate observation and gain access to the branchial cavity (link to image). The opercular or hyoid hemibranch (also known as pseudobranch), located on the dorsal internal surface of the operculum, may now be observed. Any further, more detailed examinations needed according to the aims of the necropsy should be performed during this stage, such as fresh parasitological examination of gills and gill mucus, or taking samples for histology or microbiology, which should be done as soon as possible after the fish dies. Depending on the size of the specimen, routinely take either a complete arch or a section. The first arch is usually most exposed to induced or mechanical alteration, so according to the aim of the sample, give preference either to the first or second arch in order to obtain the most adequate information on the condition of the specimen. As each arch is extracted, you will gain better access to the next one and see it more clearly. It is extremely important to avoid damaging branchial filaments while extracting arches, so it is recommended to hold the gill by the arch and cut it from the internal side. Avoid touching the filaments at all times during sampling. Although the gills are protected within the cavity, their delicate structure and exposure make them especially vulnerable. In a fresh fish, gills are bright red and shiny and should not have excess mucus or appear pale and dehydrated. Thus, the condition and conservation state of the gills allow an indirect estimation of the time that has passed since the death of the specimen (see post-mortem phenomena). Pay special attention to any alterations in colour (paleness or darkening), haemorrhage, excess mucus secretion or presence of parasites, among other alterations.    

When examining integument and fins, pay special attention to the colour of the specimen to determine whether it is normal for the species. Notice also integrity of the skin, condition and tensile strength of scales, if relevant (species with scales), amount of mucus, presence of furuncles, ulcers, nodules, cottony masses or ectoparasites (see FPA). If you need to know the age of the specimen, now is the time to take samples of scales for reading and estimating age . In some farmed fish, fins are often be absent, very eroded or only present as small appendages or stumps , which should be recorded. When the aim of the necropsy is a parasitological examination, prepare slides for microscopic inspection of the integumentary mucus and certain areas of skin, fins and orifices. For example, it is important to check the internal face of the pectoral fin insertion, which is the preferred site for some ectoparasites. To do so, scrape samples of skin for observation under optical microscope or subsequent analysis.  Place a coverslip, slide, scalpel or spatula at an angle of about 45º to the base of the fin and gently scrape the surface in cranio-caudal direction. If lesions such as furuncles or ulcers are found, samples may be needed for microbiological and histological analysis. 

Eyes are vulnerable to external injury, and may also reflect certain systemic conditions. During the examination, anophthalmia, exophthalmia, intra- or peri-ocular hemorrhage, air bubbles or opacity in the lens or cornea may be found, among other alterations. A more detailed examination requires extraction of the eye, which may be left until later during the dissection, as the eye lasts longer as a sample than other parts which require more immediate attention. To extract the eyeball, insert preferably a fine-pointed, curved scissor or a scalpel between the capsule and the eye. Use forceps to hold the eye by the conjunctiva and cut the muscles following the curve of the eye, lift the eye and cut the optic nerve to free the eyeball. Keep in isotonic saline solution for parasitological analysis or place in a fixative solution (preferably Carnoy’s) for histological examination. Samples may be taken from the eye chamber for microbiological culture, which may be useful for isolating certain bacterial agents.    

Nostrils are only examined routinely for parasitological studies in order to detect possible presence of parasites in the exudates, which must be studied by smear microscopy CommentInstill water through the front nostril using a micropipette and collect it from the back nostril. The procedure should be vigorous in order to drag possible parasites from the walls of the nasal cavity..   Depending on the type of mouth structure and size of the specimen, the mouth may be opened for macroscopic observation using a forceps (link to image). During this examination, evaluate the possible occurrence of haemorrhages, erosion, cottony masses, parasites (link to image) or structural deformations, e.g. mandibles preventing the mouth from closing effectively.   Air bubbles affecting the tongue and the roof of the mouth may also be observed, as a result of gas oversaturation (link to FPA image).    

During the general macroscopic inspection of anus and urogenital papilla, note first whether there is prolapse or inflammation. Pay attention to the possible presence of haemorrhages, petechiae, reddening or pseudo-faeces, among other conditions. If necessary, optical instruments may be used for observation, and samples may be taken for microbiological and histological examination.    

After the skeletal muscle samples have been taken, if required, a more detailed analysis of the musculature can be performed, by cutting transverse slices along the flanks using a scalpel or an appropriate knife. This procedure allows for the macroscopic examination of the consistency, integrity and colour of the musculature, as well as detecting possible parasites, nodular structures, haemorrhage, abscesses, cysts or any other abnormalities.

The pericardial cavity can be opened directly, however it is easier to open the abdominal cavity first, sequentially accessing the pericardial cavity. It is useful to have at least the head, if not the entire fish, over a paper towel, which prevents the fish from moving during the dissection, and helps by absorbing inevitable leakage of body fluids. In some occasions it can be useful to have a V shaped stand to place the fish on its back. This is especially useful for blood sampling.   A traditional approach to open the belly wall, is by cutting a small incision in front, but not too close, to the vent opening, ensuring cutting through the skin and muscle layers but not the hind gut underneath. From this incision and ideally using scissors, carefully introduce the blunt tip on the slit, cutting along the medium ventral line on a caudo-cranial direction. This section can also be done using a scalpel as practice is gained. When cutting through the pelvic and pectoral girdles, they usually offer strong resistance, especially in larger animals, and stronger scissor or pliers may be required. However, maintaining a precise middle line position may prove enough to overcome the structures as they are softer at that point. Continue the cut until reaching the isthmus, the narrow, fleshy region between the branchiostegal membranes of the opercula. When cutting with a scalpel, use a serrated or tooth forceps to hold and slightly lift the belly flap to guide the cut and avoid damaging inner organs. A metal V shape probe inserted within the cavity through the initial slit and used as a guide for the scissor or scalpel, is sometimes used until sufficient practice is gained, to facilitate the section and prevent damaging the organs, particularly the rupture of the digestive tract. An alternative way to do the ventral cut is to start cranially, by making a slit in the isthmus region and cutting in a cranio–caudal direction towards the vent, stopping just before to avoid the anus and urogenital papilla.   While opening the abdominal cavity pay special attention to the possible leakage of ascitic fluid. Take note and describe the characteristics and, if appropriate, take a sample for bacteriology.   Whichever way is used for this first cut, you can continue and cut along the isthmus to open the ventral side of the pericardial cavity, however as a preference, this can be done after completing the opening of the abdominal cavity.   The following cut is intended to open a window on the side of the fish and expose the abdominal cavity. With the help of tooth forceps lift the flank of the body wall and ideally, with a curve scissors or carefully with a scalpel, start sectioning the body wall from the cranial periphery of the vent (without touching the region) upwards and depending on species, slightly posterior, following the edge of the abdominal cavity surrounding the viscera and reaching approximately the level of the lateral line. Then continue in an oblique cranial direction following the dorsal edge of the body cavity to reach the operculum, avoiding puncturing the air bladder which lies underneath. The flap, frequently including the pectoral fin, is then removed with a last vertical cut next to the posterior edge of the opercula and reaching the cranial end of the first cut in the isthmus region.  If microbiology samples are required and in order to minimise the risk of contamination, this is the stage of the necropsy to perform the sampling, before any further spillage of liquids or contents may occur as necropsy progresses.   Once the fish is open and without removing or displacing any organ or structure, assess the condition of the peritoneum and determine if the location and the appearance of the organs and are normal for the species. Visually scan the cavities and organs for abnormalities on size, colour, odour, and for the presence of adhesions, parasites and other alterations. Check and take note of “anything that shouldn’t be there or anything that should, but is not”. Fresh sample reserved for subsequent microscopic examination in situ, must be kept moist with an isotonic solution. Excessive perivisceral fat and the developmental stage of the gonads may hinder the observation of the various organs. If this is the case, carefully remove the fat as required, except that around the pyloric caeca where the diffuse pancreas is located. When the gonads are mature this may prevent observing other organs, you may have to remove them at this stage by cutting the mesovario or the mesorquio, to enable the macroscopic observation of the rest of the organs.

The pericardial cavity is considerably smaller than the abdominal cavity. To complete opening the pericardial cavity and access the heart, the initial ventral cut should be taken further towards the cranium to open the lower wall of the cavity. The abdominal and pericardial cavities are separated by the transverse septum, which in case the heart is accessed ventrally, it will not necessarily need to be cut fully open. However, it will be stretched when the body walls are separated to check inside the pericardial cavity.   Examine the cavity for the presence of blood clots, exudates or parasites. The different chambers of the heart should have different colours and consistencies, check and register abnormalities also in position, size, brightness and possible bleeding. For the removal of the heart, hold the cranial end of the bulbus arteriosus with serrated or tooth forceps and cut cranially the ventral aorta. Pulling very gently upwards to locate the connection of the atrium with the sinus venosus, and cut the hepatic and cardinal veins right alongside the septum to free the heart. Try to avoid pulling excessively as it risks damaging or rupture the atrium. The heart can be also accessed by opening through the septum from the abdominal cavity once the viscera has been removed. See images on the species carrousel.  

The digestive tract anatomy varies widely depending on the species and may present further differences in dimensions and appearance  depending on the fish alimentary status (empty tract or very full). In general terms the digestive tract consists of an oesophagus, stomach and intestine. As described in the fish diagnosis and the anatomy sections, not all species have a distinct stomach, and the length of the intestine may widely vary. The later may be coiled in several loops.

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Check and take note of the overall appearance, and register the stomach (if present) repletion. Look for signs of haemorrhage, petechiae, inflammation, parasites, nodules or cysts on the external surface of the tract. According to the objectives of the autopsy, it may be necessary to prioritize the sampling of tissues. For instance, for histological examination, the liver, pancreas, spleen and a portion of different parts of the digestive tract may be sampled at this stage. Additionally, if bacteriology samples from kidney are required, the digestive tract can be first removed for subsequent analysis, allowing the kidney sample to be taken first (see subsequent sections). To remove the digestive tract, locate the most cranial end of the oesophagus above the transverse septum, and the opposite, at the end of the intestine, at the vent region. Hold the oesophagus with a forceps and cut free as cranially as possible. Gently pulling, remove all the tract from the cavity and cut the intestine end which may also required to be occlude by holding with haemostatic type forceps to prevent leakage of the content. Accessory glands as the liver, and the spleen and potentially the air bladder will be pulled along as the tract is removed out the cavity. Once removed, it is desirable to extend the tube on a tray of dissection or a glass plate and separate the mesenteries and the excessive fat. The above will be common practice for parasitological examination.   For the opening of the digestive tract, usually performed for parasitological assessment, it is recommended to use blunt-tipped scissors to cut open the tube longitudinally. Once open, assesses the amount of mucus and the possible presence of fluids, haemorrhage, petechiae, foreign bodies and macroscopic parasites. Enteritis and the parasitic infection by helminths are some of the most frequent findings. When deemed necessary, a sample of the food content is preserved for later analysis. Similarly, for the fresh microscopic evaluation of content or mucus smears or the intestinal wall, a small portion of the tissue of interest is extended on a slide and kept moisture until examination.

These organs belong to different systems; however, for the purpose of their examination they have in common being the soft tissues (along with the kidney and brain) that are readily accessible within the body cavity. The liver is an organ with a smooth surface, easily distinguishable by its size and location in the front part of the abdominal cavity. The colour of this organ may vary widely, both between species as within, depending on the fish sex, the feeding, the reproductive status and due to diseases. Check that the colour is within normality for that species, this must be done as soon as possible as it will change with the progress of the necropsy as vessels are sectioned and due the onset of post mortem phenomena, which may lead to erroneous conclusions. Liver is a delicate organ and can easily be damaged during dissection. Avoid compression when manipulating or sampling, by using for example sharp scalpels to take cut portions rather than scissors, which will compress the organs before effectively sectioning a portion.   Frequent alterations are the increase in the size (hepatomegaly) and changes in the colour and the brightness, usually associated with nutritional or disease problems. If samples are required for histology, bacteriology, virology or molecular testing, they should be obtained prior to any manipulation of the organ. When removing the liver, the mesenteries, veins arteries and ducts need to be sectioned to cut free the organ.  Special care is necessary to prevent damage of the gallbladder, which should remain attached to the liver and avoiding puncture, as the bile is a strong acid, can damage other tissues. Sample of the bile is sometimes performed for parasitological assessment.

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A series of cuts through the parenchyma are sometimes carried out to observe the texture and colour of the cut surfaces. At this time it is possible to observe some evidence of fatty degeneration. It might be advisable to examine the gallbladder before the liver on occasions, while still attached, by turning over the organ to make it accessible. A macroscopic examination assessing the bladder repletion and the colour and transparency of the bile can be important to evaluate nutritional status. If sampling is required, separate it from the liver by cutting the hepatic duct and place in a Petri dish for fresh microscopic examination, or place directly into a fixative for histological examination.     The spleen surface is also normally smooth and with well-defined borders. The organ is located slightly ventrally in the middle area of the abdominal cavity. When there is stomach, the spleen is found in its caudal curvature. The macroscopic examination includes an assessment of the colour, brightness, size and sharpness of its edges. As with the liver, you may want to make small cuts through the organ to check consistency of the parenchyma or sample tissue for fresh smears or imprints. A possible alteration is inflammation, with the consequent loss of sharp edges and the increase in size (splenomegaly).

 

 

 

The gonads are usually pairs bodies, located on a ventral position to the air bladder. Their appearance, size and colour vary, depending on the fish species, the sex and the state of development. Size and colour are usually recorded to assess their degree of development. To remove the gonads pull lightly to locate the connection cranially, sometimes simple pulling detaches them but it is preferable to cut rather than tear. Very mature male gonads (testicles) may easily release sperm which should be avoided. Loose sperm in the histological samples of other tissues can lead to misinterpretation. After removal, they are placed in a suitable container and reserved usually for reproductive or parasitological studies. The air bladder is located dorsally to the gut and easily to observe if it has not been deflated. The colour, shape and wall consistency varies widely among species. It can be almost transparent and delicate as in salmonids, or solid pearl white in colour and strong wall consistency on cyprinids.

Its integrity must be assessed and the possible presence of parasites and exudates of different characteristics, haemorrhage, petechiae or hyperaemia, among others. To remove separate from the digestive tract and carefully using a non sharp tool like a spatula, detach the edges from the body wall leaving the kidney behind, with the scope of dissecting the bladder without loss of gas. The bladder would remain together or not the digestive tract, depending on whether the fish is physostmous or physoclist, respectively. In the first case the pneumatic duct needs to be severed. The bladder is placed in a container and kept moist for subsequent observation, especially for the purpose of parasitological examination. Once the gonads and air bladder are removed, the kidney becomes accessible. This organ is located dorsally to the abdominal cavity, intimately attached to the spine. The peritoneum sheet must be cut to have real access to the organ. Check for observable macroscopic changes, which frequently affect the posterior (excretory) part, including increase in size, swelling, thickening, loss of brightness, vesicles and pustules. Nodular or granulomatous changes are also possible observations. The presence of the the “corpuscles of Stannius”, located in the ventral wall of the organ approximately in the middle zone, marks a transition between the excretory  and the haemopoietic portions of this organ and should not be confused with granular lesions.

If required, proceed to the sampling for histopathology or microbiological examinations. The kidney is an important organ in the detection of bacterial and viral agents as well as target by fungal agents and sensitive to water quality issues. Sometimes fresh tissue samples are checked as imprints or smears stained and analysed in situ.

 

For accessing the cranial cavity and brain the skull needs to be opened. Before attempting to cut open the hard tissues of the head, secure the fish with a cloth or paper towel to prevent sliding, as it can be dangerous when using sharp tools. Depending of the fish size, a scalpel may suffice, or a sharp knife or a saw might be required in larger specimens. If using scalpels or knifes, hold firmly the head of the fish by introducing a tooth forceps on the mouth cavity, and proceed to make a decisive sharp cut on a horizontal plane in a line just above the eye balls, continuing slightly into the dorsal musculature. The section has to aim to go through from side to side of the head in  the same cut, to open clean the roof of the skull.

Once exposed, observe in situ the cavity and the brain, its general appearance, the colour and consistency, excessive fluids, haemorrhages, as well as the eventual presence of macroscopic parasites. The killing method applied and the time elapsed since death can cause alterations which must be taken into consideration. When histology or microbiological samples are required, these must be taken prior to any further manipulation. To extract the whole brain, depending on the correctness of the dissecting plane, the tissue will have remained on one piece either on the skull cap (having severed most of the neural connections) or would be still in the cranial cavity, but now clearly visible from top. Both options allow a clean and easy collection; if on the top with immediate access as no connection to nerves or spinal cord remains; if in the cavity, after sectioning the olfactory and optical nerves that will hold to the front and the spinal cord, to the back. For the detection of protozoa, fungi and certain parasites, fresh tissue preparations are checked in situ. The open skull will also provide access to locate and remove the otoliths, which are used in the age determination, mainly in fish without scales.