Necropsy-based Wild Fish Health Assessment

Methods described here have been approved by the Leetown Science Center's Institutional Animal Care and Use Committee.

1. Fish Collection

1. Collect live fish with a minimum of stress. Use boat or backpack electrofishing, hook and line or nets.
2. Hold fish in live wells or aerated containers until sampling.
   NOTE: The American Fisheries Society has published a number of guides for fish collection, handling and anesthesia/euthanasia^26,27,28. Wear gloves when handling fish.

2. Fish Necropsy

1. Euthanize a fish.
   1. Place fish in the anesthetic until opercular movement ceases and the fish loses equilibrium. After another 2–10 min the fish will be euthanized; however, this may also vary by species.
      NOTE: Fish can be euthanized with a number of anesthetics (see Table of Materials for most commonly used). The method of euthanasia will depend on the laboratory measurements that will be conducted on tissues collected²⁹.
2. Measure biometric characteristics.
   1. Weigh the fish to the nearest gram.
   2. Measure the fish length to the nearest millimeter.
      1. Measure the total length from the tip of the snout with the mouth closed to the end of the tail when pinched together.
      2. Measure the fork length from the fork in the tail to the tip of the snout, and the standard length from the tip of the snout to the end of the body (beginning of the tail).
   3. Calculate the condition factor using the following formula:
      Condition factor = (total body weight – gonad weight)/total length³.
      Note: Gonad weight is subtracted from the total body weight since gonads can contribute significantly to the total body weight, particularly in prespawn female fish.
3. Obtain a blood sample.
   NOTE: Blood is most commonly taken from the caudal vein but can also be withdrawn from the dorsal aorta or by cardiac puncture³⁰.
   1. Extract a peripheral blood sample from the caudal vein with a 22 or 23 G needle on a 1 to 5 mL syringe, depending on size of the fish. Insert the needle anterior to the caudal area below the lateral line (Figure 1A and 1B). Angle it upwards until hitting the spine and then withdraw slightly. The vein is ventral to the overlying spine.
      NOTE: If blood smears will be made or serum is required, no anticoagulant is used. In most cases, plasma will be collected and, hence, an anticoagulant such as sodium heparin, EDTA or lithium is used to coat the needle and syringe and is also in the blood collection tube (e.g., vacutainer).
   2. Remove the needle and place into a sharps disposal container prior to putting the blood into the collection tube.
      NOTE: Blood can be held on ice but depending on subsequent analyses should be centrifuged as soon as possible³⁰.
   3. If nuclear abnormalities or differential blood counts will be evaluated, immediately place a drop of blood on duplicate clean glass microscope slides. Back a second slide at a 45° angle into the drop, which is then drawn across the surface by capillary action. Allow to air-dry³¹.
   4. Centrifuge blood at 1,500–2,500 x g for 15 min to sediment the cells. Remove plasma/serum with a sterile transfer pipet, aliquot into cryogenic vials, and store at -80 °C.

Figure 1: Obtaining a blood sample from a fish. (A) A recently euthanized fish is laid on its side and the lateral line located. (B) A needle is inserted ventral to lateral line (arrow), angled upward until needle touches the backbone. It is then slightly withdrawn, and suction initiated to withdraw blood. Please click here to view a larger version of this figure.

4. Conduct a necropsy-based health assessment on each fish.
   NOTE: A number of publications illustrating and describing lesions and abnormalities are available^32,33,34,35.
   1. Document external abnormalities including lesions on body surface and fins (Figure 2), eyes and gills (Figure 3), external parasites such as leeches (Figure 2D), grubs or trematode metacercarial cysts (Figure 2D, 3B) and gill parasites (Figure 3D). Document type, location and size of observed abnormalities on data sheets, as well as photographically, if possible.
   2. Open the abdominal cavity (Figure 4A) using a scissors by cutting from the anal area to the operculum and then removing the flap of muscle to expose the internal organs.
      NOTE: If anterior kidney will be collected for immune function (see step 5 below) or samples collected for bacteriology or virology, the external body surface should be disinfected with 70% alcohol and those samples should be obtained before the necropsy performed. If tissues are only being used for visual observations, plasma analyses and histopathology sterile technique is not necessary.
   3. Document internal abnormalities (Figure 4) including general or focal discolorations of the various organs (Figure 4B-4D), presence of raised areas (Figure 4E), cysts, parasites, and size abnormalities (enlarged, atrophied).

Figure 2: Examples of visible lesions observed on body surface and fins of fish. (A) A small, slightly eroded lesion (arrow) on the lateral body surface. (B) A large reddened area (arrow) involving the caudal body surface. (C) Raised, black lesions (arrows) on the body surface and fins. (D) Leeches (white arrow) and small black spots (black arrows) on the fin. Scale bar = 3 mm. (E) A raised, multilobed, pale lesion (arrow) on the body surface. Please click here to view a larger version of this figure.

Figure 3: Examples of visible lesions of the gills and eyes of fish. (A) A pale area (arrow) within the lens of an eye. Scale bar = 5 mm. (B) White cysts (white arrows) and small black spots (black arrows) caused by trematode parasites on the operculum covering the gills (a). Scale bar = 1 cm. (C) A pale, eroded area (arrow) on the gill (a). Scale bar = 5 mm. (D) A gill that has been removed showing parasites (arrows) attached to the gill filaments. Scale bar = 2 mm. Please click here to view a larger version of this figure.

Figure 4: Examples of a necropsy and internal abnormalities of fish. (A) During a necropsy the fish is cut open (along the white arrow) and a flap of muscle (black arrow) removed to expose the gonad (a) and the spleen, being held by forceps and scissors. (B) Mottled liver (a), testes (b), intestine surrounded by adipose fat (c) and stomach (d). Scale bar = 5 mm. (C) Liver (a) with a dark red area (arrow), ovary (b) and intestines (c). Scale bar = 5 mm. (D) Liver with greenish discolored areas (arrows). Scale bar = 1 cm. (E) Example of a normal (a) and abnormal (b) testes with raised nodules. Scale bar = 1 cm. Please click here to view a larger version of this figure.

5. Obtain hepatosomatic index (HSI).
   1. Remove liver by severing the hepatic artery and the connective tissue of the anterior end. Gently lift out while trimming adhesions and other connections to intestine and adipose fat. Take care to not puncture the gallbladder. Weigh the liver.
      NOTE: Some fish, such as cyprinids, do not have a discrete liver but rather hepatic tissue wrapped around intestines and other organs. For these species, it may not be possible to obtain liver weights.
   2. Calculate hepatosomatic index (HSI) using the formula:
      HSI=liver weight/total body weight
6. Calculate the gonadosomatic index.
   1. Remove the gonads and weigh it.
   2. Calculate gonadosomatic index (GSI) using the formula:
      GSI = gonad weight/total body weight

3. Preserve Tissues for Microscopic Pathology

NOTE: A number of fixatives including 10% neutral buffered formalin and Z-fix, a formalin-based fixative with zinc, can be used for preservation of tissue in the field. The latter is preferred if methods such as in situ hybridization or fluorescent antibody staining may be used.

1. Carefully cut but do not pull out tissue samples. Keep individual tissue pieces <2 cm in size and <5 mm thick for proper fixation. As a rule of thumb, use approximately 10x more fixative by volume than the tissue for proper preservation. Place all tissue samples from one fish in the same leak-proof container of the appropriate size, depending on the size of fish being sampled.
2. Place pieces of any external abnormalities in the fixative container. Also, include an adjacent piece of normal tissue.
   NOTE: Improper handling such as compression or other mechanical damage, long exposure to air or sunlight, and freezing can cause artifacts.
3. Cut at least five 3–4 mm thick pieces of liver from various regions and place into the fixative container. Include normal and abnormal areas, if observed.
4. Depending on size, place a whole gonad or multiple pieces along one gonad into the fixative container.
5. Place either whole organs, if small, or pieces of all other organs (spleen, anterior and posterior kidney, gills, heart, intestine and stomach) in the fixative container. If abnormal tissue is observed, preserve an adjacent piece of normal tissue as well.

4. Remove the Otoliths for Age Analyses

NOTE: Age can be an important variable in fish disease/fish health studies. While a number of structures, including scales and spines, have been used for age determination, most studies comparing structures have found the otoliths to give the best results^36,37. Teleost fishes have three pairs of otoliths – lapillus, sagitta and asteriscus. Generally, the sagittal or lapillus otoliths are collected for aging although that may vary by species. Removal and aging techniques have been previously described³⁸.

1. Cut through the gill isthmus, and bend the head back. Strip away connective and muscular tissue around the inferior portions of the neurocranium to locate prootic bullae, a raised bony area.
2. Score or cut with bone cutters and crack to expose the otoliths. They can be seen with the naked eye.
3. Place otoliths in a labelled vial or a coin envelope and store at room temperature until analyzed for age by counting the rings or increments³⁸. If placing in a vial, open cap once returned to the laboratory and allow to thoroughly dry prior to storage.

Figure 5: Removal of otoliths. (A) The isthmus is cut and the connective tissue and muscle pulled away to expose the base of spine and neurospinal area. (B) The bone is cracked to expose the otoliths. (C) Lapillar otoliths are removed. Please click here to view a larger version of this figure.

5. Obtain Tissue for Immune Function Assays

NOTE: The anterior kidney is the major hematopoietic organ, the source of lymphocytes and macrophages for functional assays, and must be removed aseptically if cells will be cultured for functional assays, such as mitogenesis, phagocytic and killing ability of macrophages^39,40.

1. Spray the external surface of the fish with 70% ethanol. Use sterile scissors, a scalpel and forceps to open the abdominal cavity and remove the anterior kidney tissue, which is a dark red organ located anterior to the swim bladder.
2. Place the anterior kidney sample in media (e.g., Leibovitz's L-15) to keep the cells alive. Homogenize kidney samples with a sterile hand-held tissue grinder (e.g., Tenbroeck tissue grinder) into single cell suspensions. Hold on wet ice until returned to the laboratory.

6. Preserve Tissue for Nucleic Acid Analyses

NOTE: If downstream molecular analysis will be conducted, such as gene expression using transcript abundance⁴¹ or quantitative PCR⁴² (polymerase chain reaction), place the pieces of tissue to be assessed in an appropriate preservative (e.g., RNAlater stabilization solution) as soon as possible.

1. For RNA preservation, place two to three small (2–3 mm) pieces in the appropriate preservative at a 10:1 ratio of preservative volume to tissue.
   NOTE: Samples should be shielded from sunlight or excessive heat and transported on wet ice.
2. For DNA preservation, place two to three small pieces of tissue into 95% ethanol (10:1 ethanol to tissue by volume). Then hold the samples on wet ice and then store at -20 °C.