- Email: [email protected]
Fish Necropsy
CHAPTER
Fish Necropsy Jeffrey P Fisher
m
Pentec Environmental, Inc., Edmonds, Washington, USA
Z
Mark S Myers
O
m
National Marine Fisheries Service, Northwest Fisheries Science Center, Seattle, Washington, USA
@ "IB
Introduction Careful necropsy technique is critical to establish reproducibility and credibility in fish anatomy (Harder, 1975; Rich and Smith, 1978; Yasutake and Wales, 1983; Roberts and Sheperd, 1986; Chiasson and Radke, 1996), pathology, toxicology, immunology, neurobiology and taxonomy. In pathology, fish necropsy is employed for the purposes of diagnosing enzootic and epizootic infection, for characterizing idiopathic or toxicopathic disease, and for satisfying requirements of routine fish health inspections required for the marketing of food fish cultured throughout many parts of the world (Strange, 1983; Phillips, 1988; Hunn and Schnick, 1990; Reimschuessel, 1993; AFS, 1994). Aquaculture research has additional needs where necropsy is required, such as studies to evaluate the effects of dietary constituents on body composition and food conversion efficiency as reflected by somatic indices such as liver-to-body-weight ratios (Brown and Gratzek, 1980; Anderson and Gutreuter, 1983). Copyright ~ 2000 Academic Press
Although this text is devoted to the use of the fish as a laboratory animal, fish are often used as sentinels of chemical contaminant exposure in the environment. As such, field necropsy may be required to investigate fish kills or to establish ecological risks from sublethal levels of toxicant exposure by assessing aspects of their internal condition. Such studies could involve tissue resection for various chemical contaminant analyses (Stehr et al., 1993; Hunn and Schnick, 1990), biopsy for histopathology (Ostrander et al., 1993; Wooster et al., 1993), or a broad assessment of somatic indices that collectively may indicate population stress when compared against unexposed populations (Adams et al., 1993). Somatic and stress indices obtainable through field necropsy are also possible with focused laboratory-based toxicological research, where, for example, necropsy may be required to identify systemic lesions resultant from acute or chronic toxicant exposure. The advantage provided by the laboratory setting is that more sophisticated techniques can be applied and more control of the conditions under which the necropsy occurs can be maintained. Neurochemistry
O r H'I
r m
and behavioral toxicology of fishes may require the dissection of the fish brain, a procedure more amenable to laboratory facilities where appropriate control over necropsy conditions can be established. Chapter 32 illustrates techniques that may be used to perform fish necropsies in fish of the salmoniform and pleuronectiform body types. The techniques can be applied to fishes of different morphology, with only minor variations. The techniques illustrated here do not assume the necessity of a sterile field, although such conditions are often required for fish disease diagnosis or surgery. (Specific procedures for those protocols are detailed elsewhere in the handbook).
1r O u
Salmoniform fish type
Z =!= 1
@ I,i,/ ev-
uA k3
O r r
Characterize the epidermal integrity and condition of all fins. Record the location, size, and severity of ulcerations, abrasion, erosion, hemorrhage, other gross lesions, and parasitism. Extend all fins for proper inspection (Figure 32.1). Inspect the dorsal and ventral surface of the specimen. Distend the pelvic, pectoral and anal fins; record fin lengths and condition, and excise with scissors or scalpel if needed for study (e.g. histopathology). Examine vent (anus) for signs of hemorrhage, discharge, and/or parasitism (Figure 32.2). Identify any ocular and dermal abnormalities and anomalies. For example, the specimen pictured exhibits periorbital hemorrhage (Figure 32.3). In a healthy fish, the lens and overlying cornea should be clear with no abrasions or discoloration. Note obstructions or abnormalities within the buccal cavity (Figure 32.4). Gill rakers should be clearly separated and fully developed. Deflect the operculum (gill cover) toward the head and make a cut with scissors along the preopercular margin, beginning at the ventral margin of the preoperculum near the base of the gill arches (Figure 32.5). Examine the pseudobranch on the ventral surface of the preoperculum (not pictured). Examine the gill filaments, gill arches and gill rakers. Record external gill architecture, noting color, erosion, necrosis, hemorrhage, exudate and typical signs of parasitism (e.g. cysts) (Figure 32.6).
Using scissors, clip the outer gill arch with attached gill filaments at the ventral and dorsal connections. Underlying gill arches can be removed sequentially. The gills pictured exhibit mild erosion along the outer margin of filaments. Make an incision immediately rostral to vent (Figure 32.7). Avoid perforation of intestine or rectum. If sterility is required (e.g. for bacteriological or virological samples), swab external surface with disinfectant such as an iodophore or alcohol prior to making incision. Use blunt-nosed scissors and cut in an anterodorsal direction through the hypaxial musculature, arcing slightly above and around the internal organs within the peritoneal cavity. Make a second anterior (rostral) cut along the ventral midline. The anterodorsal cut should eventually be below the lateral line approximately 4-6 scale widths. Avoid contact with internal organs. Reflect the hypaxial musculature overlying the body cavity, and at the dorsal opercular margin make a delicate anteroventral cut around the gills and pericardium, taking care to avoid puncturing the heart (Figure 32.8). Examine the condition of the visceral milieu (Figure 32.9). For example, the stomach should be elastic, the spleen should be dark red and firm, the liver should be deep pink to red and soft, but not friable. The pyloric ceca should be filamentous and fully extended with intact mesentery between individual ceca. Record signs of abnormalities, such as the following: size, shape and numeric anomalies; density, consistency and textural alterations; color anomalies; fluid and hemodynamic anomalies (e.g. ascites, edema, congestion, hemorrhage, hematoma); cysts and nodules; and more general anomalies such as exudate, erosion, distention, rupture, ulcerations, and grossly visible parasites. To remove the visceral mass, begin by grasping the anterior portion of the esophagus with forceps, and while gently pulling the esophagus caudally, transect the esophagus with scissors at the esophageal sphincter (Figure 32.10). Be careful to avoid perforation of the gall bladder in this step. Displace the alimentary tract and other associated viscera from the trunk cavity from the rostral position and, with scissors, transect the rectum immediately rostral to the anus (Figure 32.11). Cut and tease away mesenteric attachments (connective tissue) between the alimentary tract and liver (Figure 32.12). Extend the components of the alimentary tract for determination of architectural integrity (Figure 32.13). To harvest the spleen, clasp it by its attached mesentery and cut it away from remaining attached viscera.
I ~omHc~Nm~
a om~c,~
.....
~lll~}~l!ll!!~YVlllUJl~ll lllJl~jljl]ill~luLl:tlilil~hl I I I l l l ~ l j l i ~ 1 1 ~
"11
Z
rl'l
t~ HEAD REGION (brain and respiratory tissue)
I
TRUNK REGION (body cavity)
I
TAIL REGION (caudal to vent)
Figure 32.1 External examination: lateral view.
O .<
O "O O
N l-r1
~D C m
Figure 32.2 External examination: ventral view.
Figure 32.3 External examination: head region.
Separate mesenteric attachments connecting the gall bladder to liver and upper intestines. Clasp bile duct with forceps and cut bile duct with scissors, taking care to avoid spillage of bile from bladder on to the liver (Figure 32.14). The bile can be subsequently harvested by perforating the bladder with a No. 11 scalpel blade while suspended over the mouth of an amber glass vial (Figure 32.15). Such a procedure is effective for analyzing a variety of fluorescent aromatic compounds, primarily
representing metabolites of polycyclic aromatic hydrocarbons (Krahn et al., 1986). Cut caudally through the esophageal and stomach walls to expose the esophageal and gastric mucosa (Figure 32.16). Record the presence or absence of food, the quantity and quality of that observed, and any irregularities in the mucosal architecture (parasitism, hemorrhage, ulceration, etc.). Proceed caudally to reveal the pyloric sphincter (separating the stomach from the upper intestine), and the mucosa of the upper and lower intestine (Figure 32.16).
Figure 32.7 Internal examination: entry incision.
Q.
0e,e
~d
Figure 32.4 Buccal cavity examination.
Z Hypaxial musculature
m
LL
O
Epaxial musculature (above lateral line)
Figure 32.8 Internal examination: entry.
iii
iii
L.)
9
t-~
(3-
Figure 32.5 Gill examination: operculum removal.
.....
nyl../a^lo
i I i iu:)l.u
la i.u i ~
I]illjllilllllltll]tlll][iitLhiil'~!liti~iiiiitittilitlllll/llltillil 9
9
i~
i
;~J'
T
l
i~
t
I1[ , ~ t l l l 2 u
"
Figure 32.9 Internal examination: characterizing organ condition.
Figure 32.6 Gill examination: tissue excision.
Expose the pericardial cavity by carefully cutting through the pericardium (Figure 32.17). This procedure can be conducted either before or after the removal of the viscera. Clasp bulbus arteriosus with forceps and transect at the rostral end to remove the heart from the pericardial cavity. Examine the structural integrity of the heart chambers. The bulbus should be elastic, non-friable and cream colored. The conical-shaped ventricle should be firm and light red;
Esophagus
Start o f upper intestine
Liver
~liliil Spleen
Pyloric ceca
Gall bladder
"~ ~
.....
Lower intestine
Figure 32.13 Alimentary tract: gross inspection. Figure 32.10 Internal examination: resection of alimentary tract (rostral).
.
Stomach
-n -i-
Z m
o "u u~
.< Figure 32.14 Alimentary tract: resection of gall bladder.
Figure 32.11 Internal examination: resection of alimentary tract (caudal).
O o r'~ r-r1 ~D C :o
.
.
.
.
.
.
.
.
.
.
.
.
Figure 32.12 Alimentary tract: separation of internal organs.
the atrium and sinus venosus should be loose and saclike, with a deep, ruby red color (Figure 32.18). In smaller fish, as pictured, the sinus venosus will be indiscernible without microscopy. Note the condition of the dorsally positioned gonads (Figure 32.19). In immature fish or fish captured outside of their reproductive cycle, they may
Figure 32.15 Alimentary tract: bile harvest.
Gastric mucosa
Figure 3 2 . 1 9 Gonads (testes and ovaries): inspection and resection.
Figure 32.16 Alimentary tract: stomach and intestines inspection.
>ul Q. 0n. u u.i
Z 2: m
LL
0 LU C]C
Figure 32.17 Heart: exposure of the pericardial cavity.
CULTURES
Figure 3 2 . 2 0 Swimbladder: inspection and resection.
c~ U,.I
U 0~c
c1.
Ventricle
Figure 32.18 Heart: inspection and resection.
not be grossly visible. Color is highly variable based on maturity and species. Those pictured are immature ovaries. Gonads in mature fish may occupy a more ventral position and represent the largest component of the visceral mass. The swimbladder is dorsally situated above the visceral mass overlaying the kidney. In the Salmonidae it connects rostrally to the esophagus through a pneumatic duct (not pictured). Transect the swimbladder immediately rostral to its caudal connection (Figure 32.20). Excise the swimbladder by
reflecting the membranous sac rostrally and transecting at the pneumatic duct. Upon removal of the swimbladder, the anterior, middle and posterior kidney are clearly observed in their retroperitoneal position, immediately ventral to the spinal column. The paired opisthonephrir ducts and urinary bladder can be seen overlying the posterior kidney (Figure 32.21). To resect a kidney section (e.g. for histopathology), make a longitudinal incision through the overlying peritoneal membrane in the caudal direction. Transect the kidney tissue at the rostral end of a desired tissue block and cut dorsally down to the vertebral column. Proceed with two oblique and caudal cuts along both sides of the kidney in similar fashion and remove the underlying tissue block (Figure 32.22). Gently lift the kidney section away from the spinal column and transect it at the caudal end (Figure 32.23). To expose the brain, use a sharp scalpel to shave the overlying skin, cartilage, and bone overlying the brain beginning just caudal to the nostrils (Figure 32.24). The cerebellum (metencephalon) and optic
Urinary bladder
Opisthonephric
~..,
Figure 32.21 Kidney: examination.
9
~ .....
Figure 32.25 Brain resection: transection of optic nerves.
ephalon
"I'I -1-
Z I'll
n
0"O U~
Figure 32.22 Kidney resection: initiation.
Figure 32.23 Kidney resection: completion.
Figure 32.26 Brain resection: transection of medulla.
lobes (mesencephalon) are generally prominent in predatory fish like salmonids compared to the forebrain (telencephalon). Cut laterally through the brain case on either side of the cerebellum and optic lobes. Transect the optic and olfactory nerves (Figure 32.25) and medulla oblongata (myelencephalon) (Figure 32.26). The brain may then be lifted out of the skull. Procedures for the removal and examination of the eye are similar to that described for pleuronectiform fish (see pages 554-556).
Pleuronectiform fish type
Figure
32.24
Brain dissection:
initiation.
The necropsy procedure for a pleuronectiform fish (i.e. flatfish) is essentially the same as that described above for salmonid fishes, with some important exceptions based on the lack of bilateral symmetry in flatfishes. The procedures described below for English sole (Pleuronectes vetulus), a member of the family Pleuronectidae (right-eyed flounders), are applicable
O 0 m
c m
Dorsal f i n
Pectoral fin
D r ~ r
=h im
Ca
Figure 32.27 External examination: eyed-side view of English sole.
>. 0.
0e~ ~ u Jd
Lateral line
Dorsal
fin
Epaxial
....... ,......
Caudal
Z 3: t/I m
UL
@ L/3 LIJ
E3 I~J U
0 Q_
Figure 32.28 External examination: blind-side view.
to members of this family as well as those of the Bothidae (left-eyed flounders). Begin by inspecting the condition and color of epidermis, eyes, dermis and all extended fins, and vent, viewing the specimen from both the eyed side (Figure 32.27) and blind side (Figure 32.28), recording all pertinent information, including parasitism, as described above for the salmonid example. Note that flatfish possess paired pectoral and pelvic fins, and single elongate dorsal and ventral (anal) fins, and a typical caudal fin. The specimen shown in Figure 32.28 displays a subcutaneous Philornetra sp. nematode worm in the hypaxial musculature overlying the ovarian cavity. To examine the gills, place the flatfish with its blind side facing up, deflect the operculum toward
the head, and make a cut with scissors along the preopercular margin, beginning at the ventral margin of the preoperculum near the base of the gill arches (Figure 32.29); proceed dorsally near the preopercular-opercular junction (Figure 32.30). Examine the prominent pseudobranch on the ventral surface of the preoperculum (not pictured). Using the same method described for the salmonid, examine the gill filaments, gill arches and gill rakers; record anomalies and excise the outer gill arch (Figure 32.31). In most cases, the entire gill arch can be preserved for histology by placement into a tissue cassette and then into an appropriate fixative. To expose the organs in the peritoneal cavity, with the blind side of the fish still facing upward, use scissors to make an incision immediately caudal to the
Second gill arch (first arch removed)
Figure 32.29 External examination, head and opercular regions: preparation for gill tissue excision.
Abdominal flap (reflected)
Rectum
Liver
Lower intestine
Hypaxial musculature
Ovary (blind side lobe)
Figure 32.31 Internal examination: entry into and exposure of peritoneal cavity. l-lypaxial musculature
Auricle. atrium
mH I#l
x Z m
O "O
.< Figure 32.30 Gill examination, gross, and gross internal examination: entry incision into peritoneal cavity. vent (Figure 32.30); proceed posteriorly and dorsally along the caudal margin of the peritoneal cavity, being careful to avoid perforation of lower intestine or rectum, or damaging the underlying ovary or testis. If necessary, use sterile technique as described for the salmonid necropsy. Continue this cut through the hypaxial musculature overlying the abdomen, following a roughly semicircular pattern (Figure 32.31), sequentially following the caudal and dorsal margins of the peritoneal cavity. Make a second anterior (rostral) cut along the ventral midline between and past the bases of the pelvic fins, and finish the exposure of the visceral organs by making a cut dorsally along the anterior margin of the peritoneal cavity; reflect the main abdominal flap dorsally and rostrally (Figure 32.31). Complete this process by carefully trimming any remaining hypaxial muscle near the anterior margin of the peritoneal cavity, posterior to the gills and pericardium, and anterior to the base of the blind side pectoral fin. Avoid contact with internal organs when making these cuts, especially the final one where the pericardium can easily be punctured. Examine the condition of the visceral organs and record any anomalies (Figures 32.31 and 32.32), as in the method described for salmon.
O O m
Bulbus arteriosus
Ventricle
Rectum
Liver
Ovary (blind side lobe)
Figure 32.32 Internal examination: exposure of heart, examination of internal organs.
To expose the pericardial cavity, carefully make a scissors cut through the pectoral girdle and musculature associated with the cleithrum and coracoid bones, and through the pericardium (Figure 32.32); discard the blind-side half of the pectoral girdle. This procedure can be conducted either before or after the removal of the viscera. In this chapter, this procedure was done for the purposes of accurately displaying the main components of the heart in situ in a flatfish species (Figures 32.32 and 32.33). In practice in the field, the heart is typically collected by first cutting the exposed pericardium (just rostral to the anterior margin of the peritoneal cavity) with a scalpel blade. Next, reach anteriorly into the pericardial cavity with forceps, and clasp the anterior portion of the bulbus arteriosus with forceps and transect it at the
c m
.... ~
~ ~,+,
:*:+
Liver
Ovary
Ovarian cavity
Anal, ventral fin
Figure 32.34 Internal examination: gonads (ovary), examination and tissue collection. Gall bladder
Posterior kidney
a.
0 s
Z -r D
IJL
@ uJ
Bulbus arteriosus
Ventricle
Auricle, atrium
Figure 32.33 Internal examination: heart, examination and tissue collection.
s
LU U
0Cz: r~
rostral end to remove the heart from the pericardial cavity. Examine the condition of the heart chambers (Figure 32.33), as is done for the salmon. In most cases, the heart can be preserved whole in a tissue cassette and placed in fixative; for heart tissue samples > 3 mm in thickness, portions of the heart may have to be bisected with a scalpel blade. Gonadal tissues in adult flatfish are present bilaterally as two lobes at the posterior margin of the peritoneal cavity, and in the case of the ovary, extend posteriorly and bilaterally into two cone-shaped cavities that are extensions of the peritoneal cavity (Figure 32.34). Ovarian tissue is most easily collected by first making a cut with scissors posteriorly along the ventral margin of the entire length of the ovarian cavity on the blind side, followed by grasping the exposed posterior end of the ovary and gently pulling the ovary anteriorly. During this process, trim any fascia or connective tissue attachments between ovary and peritoneum with fine scissors, proceeding from the posterior to anterior end of the ovarian cavity
Esophagus
Liver
Ovary (eyed side lobe)
Figure 32.35 Internal examination: resection of visceral mass, transection of esophagus.
(Figure 32.34). The entire lobe of the ovary can then be removed, placed flat on a necropsy board and examined routinely. The same process can also be conducted for the eyed-side lobe of the ovary by turning the fish eyed side up and conducting the same procedure. Sections for histology can be excised as a 3 mm transverse section cut from the middle region of the conically shaped ovarian lobe. Testes (not shown) are also situated in the posterior margin of the peritoneal cavity, but do not extend into the posterior musculature as the ovaries do, and are easily excised with scissors. The visceral mass of flatfish is removed using identical methods to those for salmonid and other bilaterally symmetrical fishes, involving initial transection of the anterior esophagus (Figure 32.35) followed by transection of the rectum (Figure 32.36). Take special care to avoid perforation of the gall bladder in this
Esophagus
Stomach
Ovary (eyed side lobe)
Rectum
Liver
1
Figure 32.36 Internal examination: resection of visceral mass, transection of rectum.
Liver
Spleen
Soleen
Figure 32.38 Internal examination: resection of gall bladder for bile collection.
Lower intestine
Gall bladder
.
Posterior kidney
"!I
Upper intestine
lr
Z
rll
r O "ID
@ Figure 32.39 Internal examination: resection of posterior kidney.
or rT1
Esophagus
Stomach
Bile duct
Pyloric sphincter
Pyloric ceca
Figure 32.37 Internal examination: examination of organs of visceral mass after resection; eyed-side view.
step. After removal from the abdominal cavity, the entire visceral mass is then rotated 180 ~ along its long axis and placed on a clean, flat surface to reveal all organs and the mesenteric attachments among them (Figure 32.37). Cut and tease away mesenteric attachments between the alimentary tract and liver (Figure 32.38). Bile may be then collected from the gall bladder using the same method as described for salmonids, beginning first by separating mesenteric attachments connecting the gall bladder to liver and upper intestine (Figure 32.38). The spleen (Figure 32.38) is collected by clasping it by its attached mesentery, cutting it away from remaining attached viscera, and then bisecting it with a scalpel to view the cut surface (not shown). If the thickness of the bisected spleen is > 3 mm, it is then placed into a tissue cassette and pre-
served in fixative - otherwise it must be trimmed further. The liver is now separated from the alimentary tract and examined for any anomalies. Sections of liver for histology are cut through its entire depth from either the longitudinal or vertical axis at < 3 mm using very sharp scissors; the sections are placed in a tissue cassette and fixed routinely. Examination of the alimentary tract (e.g. stomach, Figure 32.38) is conducted exactly as described previously for salmon. The kidney in flatfish is located retroperitoneally at the dorsal margin of the peritoneal cavity (Figure 32.39) and ventral to the vertebral column; it is not separable grossly into anterior and posterior kidney. Resection of kidney tissue in flatfish (Figures 32.39 and 32.40) follows the same procedure as that for salmonids, although tissue collection is somewhat more problematic because of the difficult angle from which one must approach the tissue collection. Sections of kidney for histopathological examination should not exceed 3 mm in thickness, for optimal fixation.
C m
Preoperculum
Figure 32.40 Internal examination: posterior kidney
Eye
Operculum
Otolith
Cranial cavity
resected.
Preoperculum
Preopercularoperculum junction
Figure 32~ Collection of otoliths for fish age determination: extracting an otolith. Operculum
>. O. 0 ~d
Z 3: ~L
@ LIJ n~ U 0a=
Dorsal fin Figure 32.41 Collection of otoliths for fish age determination: lining up the incision.
Q_
In the necropsy of flatfish for research in any discipline (e.g. histopathology, toxicology, immunology, epizootiology, stock assessment), it is highly advisable to collect otoliths ('earbones') from the labyrinths or cavities lateral to the medulla oblongata, for determination offish age. There are three otoliths, each within a sac of the labyrinth organ, but in practice it is rarely possible to collect more than two. In English sole and similar flatfish species, otoliths are routinely collected by placing the fish eyed side up, and making a vertical incision (with a sharp knife or scalpel blade) at a slightly anterior angle through the depth of the cranium at a level approximately midway between the posterior edge of the preoperculum and the posterior edge of the operculum (Figure 32.41), on the dorsal side of the head. Note position of scalpel blade in Figure 32.41, which represents the position at which this cut is made. After opening the incision further by
Eye
Scissors
Cranial
Spinal
cavity
cord
Figure 32.43 Brain dissection: initial cut along dorsal margin of cranial cavity.
forcing down on either side of the cut, forceps are then used to extract one or two otoliths from the otolith organs lateral to the medulla (Figure 32.42). In practice in the field, otoliths are typically collected just after collection of routine biological information (length, weight, sex) and collection of blood for serological, immunological, or endocrinological studies. This cut also serves to sever the spinal column and effectively sacrifice the fish so that necropsy techniques can be carried out more easily and humanely. To collect the brain and associated organs (e.g. pituitary gland) from a flatfish, we have been successful in using the following method. After making the cut to collect otoliths, make another cut with blunt scissors along the dorsal margin of the brain cavity to near the base of the eye, being very careful to avoid contact with the fragile brain tissue (Figure 32.43). Make a similar and parallel cut along the ventral margin of
Skin and cartilage flap overlying cranial cavity and brain
mm Operculum
Figure 32.46 Brain examination: blind-side view. Figure 32.44 Brain dissection: second cut and removal of cranium.
"11 Ill
Z m
O
Figure 32.47 Eye resection: cutting of periorbital skin. Eye
Optic nerves
Olfactory lobes
Optic lobes
O "121 O
Cerebellum
m
Figure 32.45 Brain dissection: cutting of optic nerves and
c
removal of brain.
m
the cranial cavity, avoiding contact with the ventral surfaces of the brain (Figure 32.44). Grasp the posterior end of this cranial flap with a stout set of forceps and pull the flap upward and rostrally, while exerting downward pressure on either side of the cranium. This cut exposes the brain, and usually permits removal of the intact brain. However, additional scissor cuts along either side of the brain may be necessary to expose it sufficiently to permit intact removal. After the brain is fully exposed from a top view, gently grasp the medulla and pull the brain posteriorly to expose the tough, stringy optic nerves, which then may be cut with a sharp pair of scissors (Figure 32.45). The brain can now be removed from the cranial cavity for examination, including examination from the ventral view (Figure 32.46) to expose the pituitary. To remove the eye, cut through the periobital skin (Figure 32.47) and oculomotor muscles (Figure 32.48) around the orbit of the eye. Transect the optic nerve,
Figure 32.48 Eye resection: cutting of oculomotor muscles and optic nerve.
if not already completed from brain dissection (Figure 32.48). Invert the eye with optic nerve facing upward, support the eye laterally on both sides with the inside edges of forceps, and bisect the eye with a sharp scalpel (Figure 32.49). At a position near the optic nerve, transect eye through sclera, retina, vitreous body, and down through cornea to resect lens (Figure 32.50).
Figure 32.49 Bisection of eye: preparation for cut, optic nerve facing upward.
>.
a. 0 ~d u,i
Z
.Ira EL
@ klJ t~
klJ
k.3 0 r r~
Figure 32.50 Bisection of eye: cut through optic nerve, through sclera and down through cornea.
For histological preparations, the bisected halves of the eye should be placed in the tissue cassette, and eventually embedded, with the cut side down so that all tissue layers of the eye can be examined in one section.
References Adams, S.M., Brown, A.M. and Goede, R.W. (1993). Trans. Am. Fish. Soc. 122, 63-73. AFS (American Fisheries Society) (1994). In Blue Book. Suggested Procedures for Identification of Certain (ed. Finfish and Shellfish Pathogens, 4th edn J.C. Thoesen), pp. 1-14. AFS Fish Health Section, Bethedsda, Maryland. Anderson, R.O. and S.J. Gutreuter (1983). In Fisheries Techniques (eds L.A. Nielsen and D.L. Johnson),
pp. 283-300. American Fisheries Society, Bethesda, Maryland. Brown, E.E. and Gratzek, J.B. (1980). In Fish Farming Handbook, pp. 237-337. AVI Publishing Company, Westport, Connecticut. Chiasson, R.B. and Radke, W.J. (1966). Laboratory Anatomy of the Perch, 4th edn. Wm. C. Brown, Dubuque, Iowa. Harder, W. (1975).Anatomy of Fishes. Part II: Figures and Hates. E. Schweizerbart/sche Verlagsbuchhandlung (N~igele u. Obermiller), Germany. Hunn, J.B. and Schnick, R.A. (1990). In Field Manual for the Investigation ofFish Kills (eds F.P. Meyer and L.A. Barclay), pp. 30-40. US Fish and Wildlife Service, Resource Publication 177, Washington, DC. Krahn, M.M., Rhodes, L.D., Myers, M.S., MacLeod, and Malins, D.C. (1986). Arch. Environ. Contain. Toxicol. 15, 61-67. Ostrander, G.K., Blair, J.B., Hurst, J. and Stark, B. (1993). Aquat. Toxicol. 25, 31. Phillips, P.H. (1988). In Fish Diseases, pp. 95-99. Post Graduate Committee in Veterinary Science, University of Sydney, Australia. Reimschuessel, R. (1993). In Fish Medicine (ed. M.K. Stoskopf), pp. 160-165. W.B. Saunders Co., Harcourt, Brace, Jovanovich, Philadelphia. Rich, A.A. and Smith, L.S. (1978). Smith's Introductory Anatomy and Biology of Selected Fish and Shellfish. College of Fisheries, University of Washington, Seattle. Roberts, R.J. and Shepherd, C.J. (1986). Handbook of Trout and Salmon Diseases, 2nd edn. Fishing News Books, Farnham, Surrey, UK. Stehr, C.M., Myers, M.S. and Willis, M.L. (1993). In Sampling and Analytical Methods of the National Status and Trends Program National Benthic Surveillance and Mussel Watch Projects 1984-1992 Volume II: Comprehensive Descriptions of Complementary Measurements, pp. 63-69. NOAA Technical Memorandum NOS ORCA 71, Silver Spring, Maryland. Strange, R.J. (1983). In Fisheries Techniques (eds L.A. Nielsen and D.L. Johnson), pp. 337-346. American Fisheries Society, Bethesda, Maryland. Wooster, G.A., Hsu, H.M. and Bowser, P.R. (1993). J. Aquat. An. Health. 5, 157-164. Yasutake, W.T. and Wales, J.H. (1983). Microscopic Anatomy of Salmonids: An Atlas. US Department of the Interior, Fish and Wildlife Service, Resource Publication 150, Washington, DC.
Fish Necropsy Jeffrey P Fisher
m
Pentec Environmental, Inc., Edmonds, Washington, USA
Z
Mark S Myers
O
m
National Marine Fisheries Service, Northwest Fisheries Science Center, Seattle, Washington, USA
@ "IB
Introduction Careful necropsy technique is critical to establish reproducibility and credibility in fish anatomy (Harder, 1975; Rich and Smith, 1978; Yasutake and Wales, 1983; Roberts and Sheperd, 1986; Chiasson and Radke, 1996), pathology, toxicology, immunology, neurobiology and taxonomy. In pathology, fish necropsy is employed for the purposes of diagnosing enzootic and epizootic infection, for characterizing idiopathic or toxicopathic disease, and for satisfying requirements of routine fish health inspections required for the marketing of food fish cultured throughout many parts of the world (Strange, 1983; Phillips, 1988; Hunn and Schnick, 1990; Reimschuessel, 1993; AFS, 1994). Aquaculture research has additional needs where necropsy is required, such as studies to evaluate the effects of dietary constituents on body composition and food conversion efficiency as reflected by somatic indices such as liver-to-body-weight ratios (Brown and Gratzek, 1980; Anderson and Gutreuter, 1983). Copyright ~ 2000 Academic Press
Although this text is devoted to the use of the fish as a laboratory animal, fish are often used as sentinels of chemical contaminant exposure in the environment. As such, field necropsy may be required to investigate fish kills or to establish ecological risks from sublethal levels of toxicant exposure by assessing aspects of their internal condition. Such studies could involve tissue resection for various chemical contaminant analyses (Stehr et al., 1993; Hunn and Schnick, 1990), biopsy for histopathology (Ostrander et al., 1993; Wooster et al., 1993), or a broad assessment of somatic indices that collectively may indicate population stress when compared against unexposed populations (Adams et al., 1993). Somatic and stress indices obtainable through field necropsy are also possible with focused laboratory-based toxicological research, where, for example, necropsy may be required to identify systemic lesions resultant from acute or chronic toxicant exposure. The advantage provided by the laboratory setting is that more sophisticated techniques can be applied and more control of the conditions under which the necropsy occurs can be maintained. Neurochemistry
O r H'I
r m
and behavioral toxicology of fishes may require the dissection of the fish brain, a procedure more amenable to laboratory facilities where appropriate control over necropsy conditions can be established. Chapter 32 illustrates techniques that may be used to perform fish necropsies in fish of the salmoniform and pleuronectiform body types. The techniques can be applied to fishes of different morphology, with only minor variations. The techniques illustrated here do not assume the necessity of a sterile field, although such conditions are often required for fish disease diagnosis or surgery. (Specific procedures for those protocols are detailed elsewhere in the handbook).
1r O u
Salmoniform fish type
Z =!= 1
@ I,i,/ ev-
uA k3
O r r
Characterize the epidermal integrity and condition of all fins. Record the location, size, and severity of ulcerations, abrasion, erosion, hemorrhage, other gross lesions, and parasitism. Extend all fins for proper inspection (Figure 32.1). Inspect the dorsal and ventral surface of the specimen. Distend the pelvic, pectoral and anal fins; record fin lengths and condition, and excise with scissors or scalpel if needed for study (e.g. histopathology). Examine vent (anus) for signs of hemorrhage, discharge, and/or parasitism (Figure 32.2). Identify any ocular and dermal abnormalities and anomalies. For example, the specimen pictured exhibits periorbital hemorrhage (Figure 32.3). In a healthy fish, the lens and overlying cornea should be clear with no abrasions or discoloration. Note obstructions or abnormalities within the buccal cavity (Figure 32.4). Gill rakers should be clearly separated and fully developed. Deflect the operculum (gill cover) toward the head and make a cut with scissors along the preopercular margin, beginning at the ventral margin of the preoperculum near the base of the gill arches (Figure 32.5). Examine the pseudobranch on the ventral surface of the preoperculum (not pictured). Examine the gill filaments, gill arches and gill rakers. Record external gill architecture, noting color, erosion, necrosis, hemorrhage, exudate and typical signs of parasitism (e.g. cysts) (Figure 32.6).
Using scissors, clip the outer gill arch with attached gill filaments at the ventral and dorsal connections. Underlying gill arches can be removed sequentially. The gills pictured exhibit mild erosion along the outer margin of filaments. Make an incision immediately rostral to vent (Figure 32.7). Avoid perforation of intestine or rectum. If sterility is required (e.g. for bacteriological or virological samples), swab external surface with disinfectant such as an iodophore or alcohol prior to making incision. Use blunt-nosed scissors and cut in an anterodorsal direction through the hypaxial musculature, arcing slightly above and around the internal organs within the peritoneal cavity. Make a second anterior (rostral) cut along the ventral midline. The anterodorsal cut should eventually be below the lateral line approximately 4-6 scale widths. Avoid contact with internal organs. Reflect the hypaxial musculature overlying the body cavity, and at the dorsal opercular margin make a delicate anteroventral cut around the gills and pericardium, taking care to avoid puncturing the heart (Figure 32.8). Examine the condition of the visceral milieu (Figure 32.9). For example, the stomach should be elastic, the spleen should be dark red and firm, the liver should be deep pink to red and soft, but not friable. The pyloric ceca should be filamentous and fully extended with intact mesentery between individual ceca. Record signs of abnormalities, such as the following: size, shape and numeric anomalies; density, consistency and textural alterations; color anomalies; fluid and hemodynamic anomalies (e.g. ascites, edema, congestion, hemorrhage, hematoma); cysts and nodules; and more general anomalies such as exudate, erosion, distention, rupture, ulcerations, and grossly visible parasites. To remove the visceral mass, begin by grasping the anterior portion of the esophagus with forceps, and while gently pulling the esophagus caudally, transect the esophagus with scissors at the esophageal sphincter (Figure 32.10). Be careful to avoid perforation of the gall bladder in this step. Displace the alimentary tract and other associated viscera from the trunk cavity from the rostral position and, with scissors, transect the rectum immediately rostral to the anus (Figure 32.11). Cut and tease away mesenteric attachments (connective tissue) between the alimentary tract and liver (Figure 32.12). Extend the components of the alimentary tract for determination of architectural integrity (Figure 32.13). To harvest the spleen, clasp it by its attached mesentery and cut it away from remaining attached viscera.
I ~omHc~Nm~
a om~c,~
.....
~lll~}~l!ll!!~YVlllUJl~ll lllJl~jljl]ill~luLl:tlilil~hl I I I l l l ~ l j l i ~ 1 1 ~
"11
Z
rl'l
t~ HEAD REGION (brain and respiratory tissue)
I
TRUNK REGION (body cavity)
I
TAIL REGION (caudal to vent)
Figure 32.1 External examination: lateral view.
O .<
O "O O
N l-r1
~D C m
Figure 32.2 External examination: ventral view.
Figure 32.3 External examination: head region.
Separate mesenteric attachments connecting the gall bladder to liver and upper intestines. Clasp bile duct with forceps and cut bile duct with scissors, taking care to avoid spillage of bile from bladder on to the liver (Figure 32.14). The bile can be subsequently harvested by perforating the bladder with a No. 11 scalpel blade while suspended over the mouth of an amber glass vial (Figure 32.15). Such a procedure is effective for analyzing a variety of fluorescent aromatic compounds, primarily
representing metabolites of polycyclic aromatic hydrocarbons (Krahn et al., 1986). Cut caudally through the esophageal and stomach walls to expose the esophageal and gastric mucosa (Figure 32.16). Record the presence or absence of food, the quantity and quality of that observed, and any irregularities in the mucosal architecture (parasitism, hemorrhage, ulceration, etc.). Proceed caudally to reveal the pyloric sphincter (separating the stomach from the upper intestine), and the mucosa of the upper and lower intestine (Figure 32.16).
Figure 32.7 Internal examination: entry incision.
Q.
0e,e
~d
Figure 32.4 Buccal cavity examination.
Z Hypaxial musculature
m
LL
O
Epaxial musculature (above lateral line)
Figure 32.8 Internal examination: entry.
iii
iii
L.)
9
t-~
(3-
Figure 32.5 Gill examination: operculum removal.
.....
nyl../a^lo
i I i iu:)l.u
la i.u i ~
I]illjllilllllltll]tlll][iitLhiil'~!liti~iiiiitittilitlllll/llltillil 9
9
i~
i
;~J'
T
l
i~
t
I1[ , ~ t l l l 2 u
"
Figure 32.9 Internal examination: characterizing organ condition.
Figure 32.6 Gill examination: tissue excision.
Expose the pericardial cavity by carefully cutting through the pericardium (Figure 32.17). This procedure can be conducted either before or after the removal of the viscera. Clasp bulbus arteriosus with forceps and transect at the rostral end to remove the heart from the pericardial cavity. Examine the structural integrity of the heart chambers. The bulbus should be elastic, non-friable and cream colored. The conical-shaped ventricle should be firm and light red;
Esophagus
Start o f upper intestine
Liver
~liliil Spleen
Pyloric ceca
Gall bladder
"~ ~
.....
Lower intestine
Figure 32.13 Alimentary tract: gross inspection. Figure 32.10 Internal examination: resection of alimentary tract (rostral).
.
Stomach
-n -i-
Z m
o "u u~
.< Figure 32.14 Alimentary tract: resection of gall bladder.
Figure 32.11 Internal examination: resection of alimentary tract (caudal).
O o r'~ r-r1 ~D C :o
.
.
.
.
.
.
.
.
.
.
.
.
Figure 32.12 Alimentary tract: separation of internal organs.
the atrium and sinus venosus should be loose and saclike, with a deep, ruby red color (Figure 32.18). In smaller fish, as pictured, the sinus venosus will be indiscernible without microscopy. Note the condition of the dorsally positioned gonads (Figure 32.19). In immature fish or fish captured outside of their reproductive cycle, they may
Figure 32.15 Alimentary tract: bile harvest.
Gastric mucosa
Figure 3 2 . 1 9 Gonads (testes and ovaries): inspection and resection.
Figure 32.16 Alimentary tract: stomach and intestines inspection.
>ul Q. 0n. u u.i
Z 2: m
LL
0 LU C]C
Figure 32.17 Heart: exposure of the pericardial cavity.
CULTURES
Figure 3 2 . 2 0 Swimbladder: inspection and resection.
c~ U,.I
U 0~c
c1.
Ventricle
Figure 32.18 Heart: inspection and resection.
not be grossly visible. Color is highly variable based on maturity and species. Those pictured are immature ovaries. Gonads in mature fish may occupy a more ventral position and represent the largest component of the visceral mass. The swimbladder is dorsally situated above the visceral mass overlaying the kidney. In the Salmonidae it connects rostrally to the esophagus through a pneumatic duct (not pictured). Transect the swimbladder immediately rostral to its caudal connection (Figure 32.20). Excise the swimbladder by
reflecting the membranous sac rostrally and transecting at the pneumatic duct. Upon removal of the swimbladder, the anterior, middle and posterior kidney are clearly observed in their retroperitoneal position, immediately ventral to the spinal column. The paired opisthonephrir ducts and urinary bladder can be seen overlying the posterior kidney (Figure 32.21). To resect a kidney section (e.g. for histopathology), make a longitudinal incision through the overlying peritoneal membrane in the caudal direction. Transect the kidney tissue at the rostral end of a desired tissue block and cut dorsally down to the vertebral column. Proceed with two oblique and caudal cuts along both sides of the kidney in similar fashion and remove the underlying tissue block (Figure 32.22). Gently lift the kidney section away from the spinal column and transect it at the caudal end (Figure 32.23). To expose the brain, use a sharp scalpel to shave the overlying skin, cartilage, and bone overlying the brain beginning just caudal to the nostrils (Figure 32.24). The cerebellum (metencephalon) and optic
Urinary bladder
Opisthonephric
~..,
Figure 32.21 Kidney: examination.
9
~ .....
Figure 32.25 Brain resection: transection of optic nerves.
ephalon
"I'I -1-
Z I'll
n
0"O U~
Figure 32.22 Kidney resection: initiation.
Figure 32.23 Kidney resection: completion.
Figure 32.26 Brain resection: transection of medulla.
lobes (mesencephalon) are generally prominent in predatory fish like salmonids compared to the forebrain (telencephalon). Cut laterally through the brain case on either side of the cerebellum and optic lobes. Transect the optic and olfactory nerves (Figure 32.25) and medulla oblongata (myelencephalon) (Figure 32.26). The brain may then be lifted out of the skull. Procedures for the removal and examination of the eye are similar to that described for pleuronectiform fish (see pages 554-556).
Pleuronectiform fish type
Figure
32.24
Brain dissection:
initiation.
The necropsy procedure for a pleuronectiform fish (i.e. flatfish) is essentially the same as that described above for salmonid fishes, with some important exceptions based on the lack of bilateral symmetry in flatfishes. The procedures described below for English sole (Pleuronectes vetulus), a member of the family Pleuronectidae (right-eyed flounders), are applicable
O 0 m
c m
Dorsal f i n
Pectoral fin
D r ~ r
=h im
Ca
Figure 32.27 External examination: eyed-side view of English sole.
>. 0.
0e~ ~ u Jd
Lateral line
Dorsal
fin
Epaxial
....... ,......
Caudal
Z 3: t/I m
UL
@ L/3 LIJ
E3 I~J U
0 Q_
Figure 32.28 External examination: blind-side view.
to members of this family as well as those of the Bothidae (left-eyed flounders). Begin by inspecting the condition and color of epidermis, eyes, dermis and all extended fins, and vent, viewing the specimen from both the eyed side (Figure 32.27) and blind side (Figure 32.28), recording all pertinent information, including parasitism, as described above for the salmonid example. Note that flatfish possess paired pectoral and pelvic fins, and single elongate dorsal and ventral (anal) fins, and a typical caudal fin. The specimen shown in Figure 32.28 displays a subcutaneous Philornetra sp. nematode worm in the hypaxial musculature overlying the ovarian cavity. To examine the gills, place the flatfish with its blind side facing up, deflect the operculum toward
the head, and make a cut with scissors along the preopercular margin, beginning at the ventral margin of the preoperculum near the base of the gill arches (Figure 32.29); proceed dorsally near the preopercular-opercular junction (Figure 32.30). Examine the prominent pseudobranch on the ventral surface of the preoperculum (not pictured). Using the same method described for the salmonid, examine the gill filaments, gill arches and gill rakers; record anomalies and excise the outer gill arch (Figure 32.31). In most cases, the entire gill arch can be preserved for histology by placement into a tissue cassette and then into an appropriate fixative. To expose the organs in the peritoneal cavity, with the blind side of the fish still facing upward, use scissors to make an incision immediately caudal to the
Second gill arch (first arch removed)
Figure 32.29 External examination, head and opercular regions: preparation for gill tissue excision.
Abdominal flap (reflected)
Rectum
Liver
Lower intestine
Hypaxial musculature
Ovary (blind side lobe)
Figure 32.31 Internal examination: entry into and exposure of peritoneal cavity. l-lypaxial musculature
Auricle. atrium
mH I#l
x Z m
O "O
.< Figure 32.30 Gill examination, gross, and gross internal examination: entry incision into peritoneal cavity. vent (Figure 32.30); proceed posteriorly and dorsally along the caudal margin of the peritoneal cavity, being careful to avoid perforation of lower intestine or rectum, or damaging the underlying ovary or testis. If necessary, use sterile technique as described for the salmonid necropsy. Continue this cut through the hypaxial musculature overlying the abdomen, following a roughly semicircular pattern (Figure 32.31), sequentially following the caudal and dorsal margins of the peritoneal cavity. Make a second anterior (rostral) cut along the ventral midline between and past the bases of the pelvic fins, and finish the exposure of the visceral organs by making a cut dorsally along the anterior margin of the peritoneal cavity; reflect the main abdominal flap dorsally and rostrally (Figure 32.31). Complete this process by carefully trimming any remaining hypaxial muscle near the anterior margin of the peritoneal cavity, posterior to the gills and pericardium, and anterior to the base of the blind side pectoral fin. Avoid contact with internal organs when making these cuts, especially the final one where the pericardium can easily be punctured. Examine the condition of the visceral organs and record any anomalies (Figures 32.31 and 32.32), as in the method described for salmon.
O O m
Bulbus arteriosus
Ventricle
Rectum
Liver
Ovary (blind side lobe)
Figure 32.32 Internal examination: exposure of heart, examination of internal organs.
To expose the pericardial cavity, carefully make a scissors cut through the pectoral girdle and musculature associated with the cleithrum and coracoid bones, and through the pericardium (Figure 32.32); discard the blind-side half of the pectoral girdle. This procedure can be conducted either before or after the removal of the viscera. In this chapter, this procedure was done for the purposes of accurately displaying the main components of the heart in situ in a flatfish species (Figures 32.32 and 32.33). In practice in the field, the heart is typically collected by first cutting the exposed pericardium (just rostral to the anterior margin of the peritoneal cavity) with a scalpel blade. Next, reach anteriorly into the pericardial cavity with forceps, and clasp the anterior portion of the bulbus arteriosus with forceps and transect it at the
c m
.... ~
~ ~,+,
:*:+
Liver
Ovary
Ovarian cavity
Anal, ventral fin
Figure 32.34 Internal examination: gonads (ovary), examination and tissue collection. Gall bladder
Posterior kidney
a.
0 s
Z -r D
IJL
@ uJ
Bulbus arteriosus
Ventricle
Auricle, atrium
Figure 32.33 Internal examination: heart, examination and tissue collection.
s
LU U
0Cz: r~
rostral end to remove the heart from the pericardial cavity. Examine the condition of the heart chambers (Figure 32.33), as is done for the salmon. In most cases, the heart can be preserved whole in a tissue cassette and placed in fixative; for heart tissue samples > 3 mm in thickness, portions of the heart may have to be bisected with a scalpel blade. Gonadal tissues in adult flatfish are present bilaterally as two lobes at the posterior margin of the peritoneal cavity, and in the case of the ovary, extend posteriorly and bilaterally into two cone-shaped cavities that are extensions of the peritoneal cavity (Figure 32.34). Ovarian tissue is most easily collected by first making a cut with scissors posteriorly along the ventral margin of the entire length of the ovarian cavity on the blind side, followed by grasping the exposed posterior end of the ovary and gently pulling the ovary anteriorly. During this process, trim any fascia or connective tissue attachments between ovary and peritoneum with fine scissors, proceeding from the posterior to anterior end of the ovarian cavity
Esophagus
Liver
Ovary (eyed side lobe)
Figure 32.35 Internal examination: resection of visceral mass, transection of esophagus.
(Figure 32.34). The entire lobe of the ovary can then be removed, placed flat on a necropsy board and examined routinely. The same process can also be conducted for the eyed-side lobe of the ovary by turning the fish eyed side up and conducting the same procedure. Sections for histology can be excised as a 3 mm transverse section cut from the middle region of the conically shaped ovarian lobe. Testes (not shown) are also situated in the posterior margin of the peritoneal cavity, but do not extend into the posterior musculature as the ovaries do, and are easily excised with scissors. The visceral mass of flatfish is removed using identical methods to those for salmonid and other bilaterally symmetrical fishes, involving initial transection of the anterior esophagus (Figure 32.35) followed by transection of the rectum (Figure 32.36). Take special care to avoid perforation of the gall bladder in this
Esophagus
Stomach
Ovary (eyed side lobe)
Rectum
Liver
1
Figure 32.36 Internal examination: resection of visceral mass, transection of rectum.
Liver
Spleen
Soleen
Figure 32.38 Internal examination: resection of gall bladder for bile collection.
Lower intestine
Gall bladder
.
Posterior kidney
"!I
Upper intestine
lr
Z
rll
r O "ID
@ Figure 32.39 Internal examination: resection of posterior kidney.
or rT1
Esophagus
Stomach
Bile duct
Pyloric sphincter
Pyloric ceca
Figure 32.37 Internal examination: examination of organs of visceral mass after resection; eyed-side view.
step. After removal from the abdominal cavity, the entire visceral mass is then rotated 180 ~ along its long axis and placed on a clean, flat surface to reveal all organs and the mesenteric attachments among them (Figure 32.37). Cut and tease away mesenteric attachments between the alimentary tract and liver (Figure 32.38). Bile may be then collected from the gall bladder using the same method as described for salmonids, beginning first by separating mesenteric attachments connecting the gall bladder to liver and upper intestine (Figure 32.38). The spleen (Figure 32.38) is collected by clasping it by its attached mesentery, cutting it away from remaining attached viscera, and then bisecting it with a scalpel to view the cut surface (not shown). If the thickness of the bisected spleen is > 3 mm, it is then placed into a tissue cassette and pre-
served in fixative - otherwise it must be trimmed further. The liver is now separated from the alimentary tract and examined for any anomalies. Sections of liver for histology are cut through its entire depth from either the longitudinal or vertical axis at < 3 mm using very sharp scissors; the sections are placed in a tissue cassette and fixed routinely. Examination of the alimentary tract (e.g. stomach, Figure 32.38) is conducted exactly as described previously for salmon. The kidney in flatfish is located retroperitoneally at the dorsal margin of the peritoneal cavity (Figure 32.39) and ventral to the vertebral column; it is not separable grossly into anterior and posterior kidney. Resection of kidney tissue in flatfish (Figures 32.39 and 32.40) follows the same procedure as that for salmonids, although tissue collection is somewhat more problematic because of the difficult angle from which one must approach the tissue collection. Sections of kidney for histopathological examination should not exceed 3 mm in thickness, for optimal fixation.
C m
Preoperculum
Figure 32.40 Internal examination: posterior kidney
Eye
Operculum
Otolith
Cranial cavity
resected.
Preoperculum
Preopercularoperculum junction
Figure 32~ Collection of otoliths for fish age determination: extracting an otolith. Operculum
>. O. 0 ~d
Z 3: ~L
@ LIJ n~ U 0a=
Dorsal fin Figure 32.41 Collection of otoliths for fish age determination: lining up the incision.
Q_
In the necropsy of flatfish for research in any discipline (e.g. histopathology, toxicology, immunology, epizootiology, stock assessment), it is highly advisable to collect otoliths ('earbones') from the labyrinths or cavities lateral to the medulla oblongata, for determination offish age. There are three otoliths, each within a sac of the labyrinth organ, but in practice it is rarely possible to collect more than two. In English sole and similar flatfish species, otoliths are routinely collected by placing the fish eyed side up, and making a vertical incision (with a sharp knife or scalpel blade) at a slightly anterior angle through the depth of the cranium at a level approximately midway between the posterior edge of the preoperculum and the posterior edge of the operculum (Figure 32.41), on the dorsal side of the head. Note position of scalpel blade in Figure 32.41, which represents the position at which this cut is made. After opening the incision further by
Eye
Scissors
Cranial
Spinal
cavity
cord
Figure 32.43 Brain dissection: initial cut along dorsal margin of cranial cavity.
forcing down on either side of the cut, forceps are then used to extract one or two otoliths from the otolith organs lateral to the medulla (Figure 32.42). In practice in the field, otoliths are typically collected just after collection of routine biological information (length, weight, sex) and collection of blood for serological, immunological, or endocrinological studies. This cut also serves to sever the spinal column and effectively sacrifice the fish so that necropsy techniques can be carried out more easily and humanely. To collect the brain and associated organs (e.g. pituitary gland) from a flatfish, we have been successful in using the following method. After making the cut to collect otoliths, make another cut with blunt scissors along the dorsal margin of the brain cavity to near the base of the eye, being very careful to avoid contact with the fragile brain tissue (Figure 32.43). Make a similar and parallel cut along the ventral margin of
Skin and cartilage flap overlying cranial cavity and brain
mm Operculum
Figure 32.46 Brain examination: blind-side view. Figure 32.44 Brain dissection: second cut and removal of cranium.
"11 Ill
Z m
O
Figure 32.47 Eye resection: cutting of periorbital skin. Eye
Optic nerves
Olfactory lobes
Optic lobes
O "121 O
Cerebellum
m
Figure 32.45 Brain dissection: cutting of optic nerves and
c
removal of brain.
m
the cranial cavity, avoiding contact with the ventral surfaces of the brain (Figure 32.44). Grasp the posterior end of this cranial flap with a stout set of forceps and pull the flap upward and rostrally, while exerting downward pressure on either side of the cranium. This cut exposes the brain, and usually permits removal of the intact brain. However, additional scissor cuts along either side of the brain may be necessary to expose it sufficiently to permit intact removal. After the brain is fully exposed from a top view, gently grasp the medulla and pull the brain posteriorly to expose the tough, stringy optic nerves, which then may be cut with a sharp pair of scissors (Figure 32.45). The brain can now be removed from the cranial cavity for examination, including examination from the ventral view (Figure 32.46) to expose the pituitary. To remove the eye, cut through the periobital skin (Figure 32.47) and oculomotor muscles (Figure 32.48) around the orbit of the eye. Transect the optic nerve,
Figure 32.48 Eye resection: cutting of oculomotor muscles and optic nerve.
if not already completed from brain dissection (Figure 32.48). Invert the eye with optic nerve facing upward, support the eye laterally on both sides with the inside edges of forceps, and bisect the eye with a sharp scalpel (Figure 32.49). At a position near the optic nerve, transect eye through sclera, retina, vitreous body, and down through cornea to resect lens (Figure 32.50).
Figure 32.49 Bisection of eye: preparation for cut, optic nerve facing upward.
>.
a. 0 ~d u,i
Z
.Ira EL
@ klJ t~
klJ
k.3 0 r r~
Figure 32.50 Bisection of eye: cut through optic nerve, through sclera and down through cornea.
For histological preparations, the bisected halves of the eye should be placed in the tissue cassette, and eventually embedded, with the cut side down so that all tissue layers of the eye can be examined in one section.
References Adams, S.M., Brown, A.M. and Goede, R.W. (1993). Trans. Am. Fish. Soc. 122, 63-73. AFS (American Fisheries Society) (1994). In Blue Book. Suggested Procedures for Identification of Certain (ed. Finfish and Shellfish Pathogens, 4th edn J.C. Thoesen), pp. 1-14. AFS Fish Health Section, Bethedsda, Maryland. Anderson, R.O. and S.J. Gutreuter (1983). In Fisheries Techniques (eds L.A. Nielsen and D.L. Johnson),
pp. 283-300. American Fisheries Society, Bethesda, Maryland. Brown, E.E. and Gratzek, J.B. (1980). In Fish Farming Handbook, pp. 237-337. AVI Publishing Company, Westport, Connecticut. Chiasson, R.B. and Radke, W.J. (1966). Laboratory Anatomy of the Perch, 4th edn. Wm. C. Brown, Dubuque, Iowa. Harder, W. (1975).Anatomy of Fishes. Part II: Figures and Hates. E. Schweizerbart/sche Verlagsbuchhandlung (N~igele u. Obermiller), Germany. Hunn, J.B. and Schnick, R.A. (1990). In Field Manual for the Investigation ofFish Kills (eds F.P. Meyer and L.A. Barclay), pp. 30-40. US Fish and Wildlife Service, Resource Publication 177, Washington, DC. Krahn, M.M., Rhodes, L.D., Myers, M.S., MacLeod, and Malins, D.C. (1986). Arch. Environ. Contain. Toxicol. 15, 61-67. Ostrander, G.K., Blair, J.B., Hurst, J. and Stark, B. (1993). Aquat. Toxicol. 25, 31. Phillips, P.H. (1988). In Fish Diseases, pp. 95-99. Post Graduate Committee in Veterinary Science, University of Sydney, Australia. Reimschuessel, R. (1993). In Fish Medicine (ed. M.K. Stoskopf), pp. 160-165. W.B. Saunders Co., Harcourt, Brace, Jovanovich, Philadelphia. Rich, A.A. and Smith, L.S. (1978). Smith's Introductory Anatomy and Biology of Selected Fish and Shellfish. College of Fisheries, University of Washington, Seattle. Roberts, R.J. and Shepherd, C.J. (1986). Handbook of Trout and Salmon Diseases, 2nd edn. Fishing News Books, Farnham, Surrey, UK. Stehr, C.M., Myers, M.S. and Willis, M.L. (1993). In Sampling and Analytical Methods of the National Status and Trends Program National Benthic Surveillance and Mussel Watch Projects 1984-1992 Volume II: Comprehensive Descriptions of Complementary Measurements, pp. 63-69. NOAA Technical Memorandum NOS ORCA 71, Silver Spring, Maryland. Strange, R.J. (1983). In Fisheries Techniques (eds L.A. Nielsen and D.L. Johnson), pp. 337-346. American Fisheries Society, Bethesda, Maryland. Wooster, G.A., Hsu, H.M. and Bowser, P.R. (1993). J. Aquat. An. Health. 5, 157-164. Yasutake, W.T. and Wales, J.H. (1983). Microscopic Anatomy of Salmonids: An Atlas. US Department of the Interior, Fish and Wildlife Service, Resource Publication 150, Washington, DC.