Leslie A. Dierauf, V.M.D. 

August 1994 

U.S. Department of Commerce 

National Oceanic and Atmospheric Adrninistratiqn 

National Marine Fisheries Service 
PINNIPEBB FORENSIC, NECWOPSY AND 

TISSUE COLLECTION GUIDE 

Leslie A. Dierauf, V.M.D. 

Edited by: Dean Wilkinson and Cindy Driscoil, D.V.M 

NOAA Technical Memorandum 

NMFS-OPR-94-3 

August 1994 

U.S. Department of Commerce 

Ronald H. Brown, Secretary 

National Oceanic and Atmospheric Administration 

D. James Baker, Under Secretary for Oceans and Atmosphere 

National Marine Fisheries Service 

Rolland A. Schmitten, Assistant Administrator for Fisheries 
The National Marine Fisheries Service (NMFS) does not approve, recommend or 

endorse any proprietary product mentioned in this publication. 4 o reference shall 

be made to NMFS or to this publication in any advertising or bales promotion 

which would indicate or imply that NMFS approves, recommends or endorses any 

proprietary product mentioned herein, or which has as its purpbse an intent to 

cause, directly or indirectly, the advertised product to be used or pdrchased because 

of this NMFS publication. 
PINNIPED NECROPSY AND 

TISSUE COLLECTION GUIDE 

TABLE OF CONTENTS 

Introduction 

Legal Authority For Pinniped Handling and Care 

Pinniped Identification 

General Principles 

Photography 

Data pharacterization 

Levels 

Codes 

Purposes of Tissue Collection 

Clinical Pathology 

Blood 

Blood Collection Equipment 

Blood Collection Tubes 

Blood Collection Techniques in Pinnipeds 

Biopsies 

Skin and/or Fat 

Liver 

Bone Marrow 

Miscellaneous 

Life History Information 

Pertinent Information 

Pinniped Standard Measurements 

Standard Necropsy Procedures in Pinnipeds 

Gross Necropsy 

Equipment Necessary 

Standard Necropsy Form 

In General 

General External Examination 

Descriptors For Use During Necropsy Reporting 

Describing 

Anatomical Locations 

Size 

Color 

Shape and Form 

Consistency 

Odor 

External Examination of the Carcass 
First 

Haircoa t 

Body Orifices 

Peripheral Lymph Nodes 

Mammary Glands 

Scrotum 

Internal Examination of the Carcass 

Positioning the Carcass 

Opening the Abdominal Cavity 

General Conditions and Observations 

Stomach 

Small Intestine 

Large Intestine 

Mesenteric Lymph Nodes 

Pancreas and Pancreatic Ducts 

Spleen 

Liver 

Gall Bladder and Bile Duct 

Adrenal Glands 

Kidneys 

Ureters 

Bladder and Urethra 

Reproductive Organs 

Female 

Ovaries and Oviducts 

Uterus and Cervix 

Vagina and Vestibule 

Male 

Testes and Vas Deferens 

Prostate 

Penis 

The Thoracic Cavity 

Diaphragm 

Ribs 

The Neck 

Thyroids and Parathyroids 

Esophagus 

Mediastinal Lymph Nodes 

Thymus and Brown Fat 

The Pluck 

Trachea and Mainstem Bronchi 

Lungs and Bronchiolar Lymph Nodes 

Heart 

The Nervous System 
COLLECTION, PREPARATION AND PRESERVATION 

OF TISSUES AND SPECIMENS 

His topathology 

Additional Equipment Recommended 

Collection and Preparation of Tissues 

Preservation of Tissues 

Checklist of Histopathologic Specimens 

Parasitology 

Additional Equipment Recommended 

Collection, Preparation and Preservation of Specimens 

Bacteriology 

Additional Equipment Recommended 

Collection and Preparation of Tissues 

Preservation 

Immunology 

Additional Equipment Recommended 

Collection, Preparation and Preservation of Tissues 

Mycology 

Additional Equipment Recommended 

Collection and Preparation of Tissues 

Preservation 

Virology 

Additional Equipment Recommended 

Collection, Preparation and Preservation of Tissues 

Toxicology 

Biotoxins 

Additional Equipment Recommended 

Collection and Preparation of Tissues 

Heavy haeta1 and Anthropogenic Organics 

Additional Equipment Recommended 

Collection and Preparation of Tissues 

Shipping and Laboratory Analytical Techniques 

Miscellaneous Diagnostic Specimens 

Isoenz ymes 

Cytochrome P-450 

DNA Studies 

Tooth/ Bone/Skeletal Preparation 

Additional Equipment Recommended 

Collection and Preparation of Skeletal Remains 

Tooth Preparation for Aging Pinnipeds 
67 SHIPPING/TRANSPORT 

67 Packing Materials 

67 Shipment of Clinical Pathological Specimens 

68 Shipment of Histopathological Specimens 

68 Shipment of Microbiological Specimens 

68 Shipment of Parasitological Specimens 

68 Shipment of Toxicological Specimens 

69 Conclusions 

Appendices 

72 Appendix A: Formulations for Commonly Used Fixatives 

73 Appendix B: Laboratories Accredited by the American Association 

of Veterinary Diagnostic Laboratories 

74 Appendix C: Sample Transfer Record 

75 Appendix D: Sample Chain of Custody Form 

77 References 

FIGURES AND TABLES 

Ew 

Figure 1: 

Figure 2: 

Figure 3: 

Figure 4: 

Figure 5: 

Figure 6: 

Figure 7: 

Figure 8: 

Figure 9: 

Figure 10: 

Figure 11: 

Figure 12: 

Figure 13: 

Otariid Blood Collection Site 

Phocid Blood Collection Site 

Hindflipper Blood Collection Site 

Liver Biopsy Site 

Sexing Pinnipeds 

Pinniped Standard Measurements 

Anatomical Descriptors 

Making the Initial Incisions in a Pinniped Necropsy 

Measuring the Blubber Layer Thickness 

Tracheal Bifurcation in Pinnipeds 

The Skull Locations to Cut for Removal qf the Brain 

Major Anatomical Structures of the Pinniped Brain 

Piecing the Puzzle Together 

5 Chart 1: Blood Collection Tubes 

16 Table I: Pinniped Standard Measurements 

19 Chart 2: Sample Label 

21-23 Table 2: Standard Necropsy Form 

26 Table 3: Decision Matrix for Pinniped Tissue Collection 

52 Chart 3: Specimen Checklist for Histopathology 
INTRODUCTION 

The Oxford Old English Dictionary defines pinnilped as "pinniU-- pertaining 

to wings or fins or feathers, and "ped as footed; or -- one who "flies through the 

water, as if on wings". 

The term forensic comes from the Latin word forum and means -- pertaining 

to, connected with, or used in courts of law; or -- suitable or analogous to pleadings 

in court. 

Guide is defined as "one who shows the way, especially to a beginner, novice 

or traveler in a strange country", or something that guides. 

This Pinniped Forensic Guide is a book of instruction and information for 

beginners, novices or travelers in a new land, a guide when working with those 

marine mammals classified as pinnipeds. The following instruction on collection, 

preparation, preservation and shipment of pinniped tissues and other specimens, 

will help provide the most trustworthy data for potential use as evidence in a court 

of law. 

This manual's intent is to assist stranding network volunteers (with little or 

no formal scientific training), basic scientists (with a general knowledge of 

mammalian physiology) and professional marine mammal experts in performing 

simple and sophisticated specimen collection procedures. Whether ydur 

responsibility for pinnipeds is on the beach, in the postmortem room, pr in an 

academic or commercial laboratory facility, the procedures and metho~ologies 

discussed in this manual should assist you in your efforts. The final goal is to 

generate supportive scientific data for diagnosing the cause or causes df stranding 

and unusual mortality events involving pinnipeds. 

LEGAL AUTHORITY FOR PINNIPED HAN-DLING AND CARE 

In the United States, legal authority for working with pinnipedg is shared 

among the Departments of Commerce, Interior and Agriculture. The U.S. 

Department of Agriculture's Animal and Plant Health Inspection Service (APHIS) 

concentrates on animal welfare and humane care and transport of pirinipeds under 

the Animal Welfare Act (AWA; 7 USC 2131-2155). The U.S. Department of the 

Interior's Fish and Wildlife Service (FWS) implements management ractions with 

regard to walrus. The U.S. Department of Commerce's National Maqine Fisheries 

Service (NMFS) is charged with primary responsibilities for all other pinniped 

species under the Marine Mammal Protection Act of 1972 (MMPA; 16 USC 1361- 

1421h). 

In order for any person to "take" a pinniped from U.S. waters ar shores, that 

person, institution or commercial business must apply for and be issued a permit or 

letter of authorization (LOA) from the NMFS's Office of Protected Resources. Each 
permit or LOA allows the taking of a pinniped for a specific purpose, such as rescue 

and rehabilitation, the protection of the public, scientific research or public display. 

The actual definition of "take" as stated in the MMPA is "to hunt, capture, kill, 

harass, or attempt to hunt, capture, kill or harass". Without a permit or LOA, the 

MMPA specifically states that capturing or harassing pinnipeds or collecting 

pinniped body parts is prohibited. 

PINNIPED IDENTIFICATION 12 

All walrus, sea lions, fur seals and seals fall into the Order Carnivora, 

Suborder Pinnipedia. Pinnipeds are amphibious, air breathing mammals that 

divide their time between waters and lands of the marine environment. Their 

bodies are streamlined, and their appendages are shortened and flattened. Pinniped 

hindlimbs are paddle-like with webbed distal appendages. There are three families 

of pinnipeds, the odobenids (walrus), the otariids (the "eared" sea lions and fur 

seals) and the phocids (the "earless" or true seals). 

GENERAL PRINCIPLES 

PHOTOGRAPHY: 

In the necropsy room, a good quality camera with a 50 mm lens and an 

additional macro lens for close-up shots is essential. Use the camera, to document 

genus and species (if unsure), to catalog unusual lesions or organs dijscovered 

during the course of the examination, and to augment any descriptiops made about 

particular conditions, that may be of scientific or legal interest. Incluge any 

photographic documentation in the pinnipedls permanent hospital/~ecropsy file. 

A video camera can also be used to document specific observations during a 

necropsy procedure, but a video record cannot entirely substitute for dear, detailed 

photographic documentation. 

DATA CHARACTERIZATION (Levels and Codes)3,4: 

Data collection on pinnipeds can be prioritized into 3 categories: 

Level A, basic minimum data, including names and addresses of principle in 

stranding event, date, exact location of event, sex(es), length and condition of the 

animal(s); 

Level B, supplementary on-site data, including basic life history and slpecific event 

data, such as weather, carcass orientation, animal and human activities in the area, 

collection of parts for age determination, etc.; and, 
Level C, necropsy examination and parasite collection, including results of careful 

external and internal examination of the animal, tissues and/or specimens 

collected, preserved and shipped/transported to other facilities. 

In addition, tissue/specimen quality is categorized by Code: 

Code 1, the live animal; 

Code 2, the freshly dead ("carcass in good condition") animal; 

Code 3, the decomposed ("carcass in fair condition") animal; 

Code 4, the severely decomposed ("carcass in poor condition") animal; and, 

Code 5, the mummified or purely skeletal remains animal. 

PURPOSES OF TISSUE COLLECTION: 

For the science of pinniped medicine and pathology to advance, the 

importance of complete record-keeping and standardized operating procedures 

cannot be overemphasized. The quality of data collected and collated is directly 

dependent upon the quality control procedures and methodologies, whether on the 

beach, in the necropsy room, in the clinical laboratory or in the compater laboratory. 

It is the live animal that is the most valuable for collection of materials that 

might yield a rapid diagnosis regarding ongoing disease processes. Uhfortunately in 

the case of marine mammals, the live animal is found least often; tharefore, it is 

imperative, if given the opportunity, that the most skilled and best trained 

personnel collect data from the live pinniped, ideally a veterinarian. 

It is crucial then, that live (Code I), rescued pinnipeds undergo complete 

physical examinations, blood collections for complete blood counts and total 

chemistry profiles, and fecal examinations for the presence of parasit* or parasitic 

ova. If time and the animals' conditions allow, further diagnostics--iocluding, but 

not limited to, additional blood chemistry and serology studies, radiography, 

ultrasonography, electrocardiography, urinalysis, nasal/tooth/ throat/ranorectal 

microbiological swab cultures and sensitivities, endocrine function tests and tissue 

biopsies--should be performed to allow for the most complete data collection and 

the most accurate causative diagnoses. Live pinnipeds, rescued and rehabilitated, 

should be appropriately tagged, released, and monitored following release. 
CLINICAL PATHOLOGY 

BLOOD 

BLOOD COLLECTION EQUIPMENT: 

Collection of blood for clinical pathological examination in pinnipeds 

normally requires appropriate restraint of the animal. Handling of a pinniped, 

particularly an adult pinniped, can be difficult, and it is best to have all necessary 

equipment pre-assembled and close at hand prior to restraining the animal. The 

equipment should include: 

1. mechanical restraint equipmentsh; 

2. chemical restraint equipment7; 

3. emergency medical supplies ("crash cart"); 

4. needles, of varying lengths and gauges; 

5. syringes, of varying sizes; 

6. betadine or nolvasan solution in spray bottle or as swabs; 

7. blood collection tubes of all types (see below); 

8. microscope slides and coverslips; 

9. Diff-Quika or new methylene blue stain; 

10. microscope 

11. indelible marker pen or pencil for labels; 

12. laboratory forms for submission of samples; and 

13. hospital record to note any observations, emergency situations or other 

observations during blood collection. 

a DiffQuik, Harleco, Gibbstown, NJ 
BLOOD COLLECTION TUBES8: 

Following is a listing 

Color-Coded Tov 

RTT, red top tube 

LTT, lavender top tube 

Grn, green top tube 

GTT, gray top tube 

BTT, blue top tube 

BlTT, black top tube 

BWTT: black & white 

commonlv used blood , 

Chemical in Tube 

I none 

1 EDTA 

1 heparin 

sodium fluoride, 

potassium oxalate 

sodium citrate 

soybean thrombin, 

snake venom / Michel's media 

collection tubes: 

Use This Tube For 

serum chemistries 

serum iron 

sterile collection vial 

complete blood count 

blood typing 

serum chemistries, 

ammonia, parasites, 

amino acids, chromosomes, sulfanilates 

glucose, activated 

clotting time 

clotting factors 

(including PT, PTT) 

fibrin degradation 

products 

viral pathology 

BLOOD COLLECTION TECHNIQUES3,4: 

When collecting blood from pinnipeds by any of the techniques discussed 

below, follow these general rules. Use sterile, standard gauge needles, syringes, and 

collection tubes. During aspiration, try not to put the blood coming into the 

collection system under pressure (do not pull back hard or fast on the syringe barrel; 

use a butterfly catheter collection system if collapsing vessels are a prablem). 

After collection, if serum is desired, let the sample in the red to tube sit in an 

upright position for 15 to 20 minutes so that the clot retracts. Never I' eave the 

serum sitting above the clot longer than an hour. Once the clot is retracted, 

centrifuge the specimen (making sure to balance the blood tube with a "blank" tube 

filled with an equal amount of water) for 5-10 minutes. Remove the specimen 

tube, and, using a clean pipette or syringe, gently aspirate the serum sitting on top of 

the red blood cells. Place the serum in a new red top tube, and label it for transport 

to the laboratory. 
Send the sample to the laboratory immediately; if this cannot be done, keep 

the specimen refrigerated (at 4' C). Serum not sent to a laboratory can be stored 

frozen (at -20' C) for possible future use. 

Blood deteriorates rapidly following death. Results from blood collection in 

dead animals (those characterized as anything other than Code 1) are questionable at 

best. If blood collection is performed at all on a dead animal, it must be done 

immediately after death to be of any value at all. Serum from a recently dead 

animal (no more than 4 or 5 hours)g can be used for bacterial or viral serological 

testing; the best collection site is often the atrium of the heart or the posterior vena 

cava. Post-mortem serum should not be used for electrolyte or enzyme values, 

since these values are immediately changed at the time of death, and can be 

misleading. If a serum sample is desired from a live pinniped, collect blood, as 

described above, let the specimen sit at room temperature for 10-15 minutes, then 

centrifuge in a standard blood chemistry centrifuge for an additional 5-10 minutes. 

Then with sterile needle and syringe aspirate the serum off the top and place it into 

a sterile red top tube. 

Body fluids, cerebrospinal fluid, and joint fluid can also be collected up to 4 to 

5 hours post-mortem; place these types of specimens in sterile red top or purple top 

tubes. 

Needle lengths vary, depending on the size pinniped from which blood is 

being collected. In a juvenile pinniped, use a 1 inch, 18 gauge needle. In an adult 

pinniped, use a 3.5 inch, 18 gauge needle. 

There are 3 typical blood sampling sites for otariids and odobemids : 

-- 1. the caudal gluteal - vein -- 

The caudal gluteal vein allows collection of the largest volume of blood with 

the least danger to the technician. Locate this vein by restraining the otariid or 

odobenid symmetrically in ventral recumbency (the animal is on its belly with its 

foreflippers tucked under it, and its hindflippers straight out behind). Palpate three 

landmarks: (1) the tail base (where the tail joins the body), (2) the femoral 

trochanters (top outside portion of the most dorsal hindflipper bone), and (3) the 

dorsal midline (a line drawn centrally along the spine). Draw three imaginary lines: 

(1) perpendicular to the tail at the tailbase, (2) perpendicular to the dqrsal spine 

between the femoral trochanters, and (3) 1/3 of the distance from the femoral 

trochanters to the tailbase and perpendicular to the dorsal midline. Insert the 

needle (with appropriate size syringe attached) 1/2 inch to 1 inch to the right or left 

of dorsal midline along line (3) above [Figure 111. Position the needle perpendicular 

to the skin's surface. Pop the needle through the skin, penetrating to the hub of the 
I site 

Gluteal Vein in Otarii .......................... 

tb = tailbase 

ft = femoral trochanters 

dm = dorsal midline 

I = blood collection site 
needle. Then slowly pull up, aspirating as you do so, until the caudal gluteal vein is 

entered. The vein is not large, but can be located in this fashion if estimates are 

accurate and needle lengths are appropriate. 

-- 2. the interdivital - veins of the hindflipper - 

The interdigital veins in the webbing of each hindflipper are not particularly 

large and can often be constricted in cool temperatures. To dilate the veins, either 

place the animal in a heated room with careful supervision, or, use a heating 

apparatus, such as an athletic heat pack, or hot compresses on the webbed surface of 

the hindflipper. Use the interdigital veins only if the animal appears healthy. Once 

the blood sample is obtained, apply firm pressure for at least a minute or two to the 

affected interdigital area to prevent hematoma formation. 

-- 3. the precaval vein -- 

This venipuncture site is used when blood is essential and no other site is 

available for use. The precaval vein is located at the heart base on the right side of 

the otariid. Palpate the sternum (the breast plate), the trachea and the ribs on the left 

and right sides of the body. At the level of ribs 3 to 5, just to the right of the animal's 

sternum, insert the needle perpendicularly (attached to the appropriate size syringe) 

and aspirate to obtain the blood sample. 

There are 2 common blood collection sites in phocids : 

-- 1. the extradural intravertebral vein -- 

The extradural intravertebral vein is relatively easy to find and produces an 

adequate blood sample volume in phocids. Restrain the seal symmeti.ically in 

ventral recumbency (the animal lying on its belly with its hindflippers straight out 

behind). Palpate for the dorsal spines of the thoracic and lumbar vertkbrae, until 

lumbar vertebrae 3 and 4 are located. Insert an 18 gauge needle with a~ttached 

syringe between the two vertebral bodies, perpendicularly and slowly, but 

continuously, to the needle's hub. If the needle hits bone, walk the needle cranially 

or caudally off the bone until the intravertebral space is entered. Once in the 

extradural venous sinus, aspirate the blood sample desired [Figure 21. 
Figure 2 : The Extradural 

[ntravertebral Vein in Phocids ....................... ----------------------- 

KEY: 

dm = dorsal midline 

ft = femoral trochanters 

tv = thoracic vertebrae 

lv = lumbar vertebrae 

= blood collection site 
-- 2. the ventral (plantar) interdinital veins of the hindflipper -- 

The ventral interdigital veins are located similarly to those in the otariid; 

however, the diameter of these phocid veins is smaller. Over the second digit, or 

just to the medial side of the fourth digit (as the hindflipper sits, straight out behind 

the animal, digit one is the most medial of the digits on the hind-flipper, digit five is 

the most lateral [Figure 3]), at the trailing edge of the interdigital webbing, insert the 

needle (with syringe), almost parallel to the skin at about a 10 to 20" angle. The 

blood collected here will be an arterial-venous mixture, so be certain to apply 

pressure to the area after blood collection is complete to avoid a hematoma from 

forming. 

Any live pinniped undergoing a biopsy must be properly restrainied with 

appropriate infiltration of local anesthetic agent at the biopsy site. Only Trained 

Veterinarians Should Perform Invasive Procedures. In pinnipeds, before 

attempting any biopsy, it may be necessary to remove the hair overlying the area of 

concern. In any case, the area of concern should be surgically prepped using a 

betadine or nolvasan scrub and 70% alcohol, prior to initiating any biopsy 

procedure. Proper handling of tissues for biopsy includes avoiding crushing, rough 

handling or cauterizing. If shrinkage is expected (especially with skin or muscle), it 

is best to remove the biopsy specimen and gently lay it in its normal configuration 

on a hard piece of material, such as a piece of cardboard or a popsicle stick, before 

immersing in fixative. Once full fixation has occurred (3 to 4 days) the tissue may be 

gently teased off the hard material and left by itself in the formalin. 
SKIN and/or FAT BIOPSY: 

A skin biopsy is the simplest biopsy to perform, should such a sample be 

needed for diagnostics. Skin biopsies generally are used to diagnose dermatologic 

disease or are used for genetic life history determinations. Whenever collecting a 

skin specimen, be certain to include both normal and abnormal tissue in your 

sample. Two methods are routinely used: 

(1) Using forceps and scalpel, excise (cut out) about a I cm2 piece of skin tissue. 

Fat or blubber may also be removed at this time. Close the incision using one or two 

interrupted, absorbable sutures. If fat or blubber is being collected for toxicologic or 

contaminant analysis, for standardization, the sample should be collected from 

under the skin overlying and just cranial to the sternum's xiphoid process along the 

ventral midline of the pinniped. In the live animal, sample size will necessarily be 

smaller than that in the dead animal. 

(2) Using a commercially available biopsy punchb, made in various bore 

diameters (4 mm is a commonly used size), place the punch in contact with the skin 

and use slight pressure and a rotating motion to obtain a core of skin and blubber. 

Once obtained, the core is grasped with forceps and elevated, and any connective 

tissue severed to remove the "plug" of tissue. The skin is closed with one or two 

interrupted sutures. 

If the biopsy is being taken for histopathology, the punch or needle sample 

should be teased into a container of 10% formalin for fixation (For further detail on 

fixation and fixation solutions see Appendix A, page 72). 

LIVER BIOPSY: 

The veterinarian may choose to perform a liver biopsy if any od the following 

indications exist during initial physical examination and diagnostic work-up: 

jaundice/icterus, elevated liver enzyme levels, elevated bilirubin levels, or a history 

of exposure to toxicologic, biologic or other hepatotoxic agents. Biopsy of the 

pimiped liver is performed with the live animal under general anes&esia and 

using a commercially available biopsy needle. The preferred needle is a 5.25 inch, 

16-gauge Manghini liver biopsy needlec. To collect an adequate liver $iopsy, the 

pinniped must be chemically restrained in left lateral recumbency (with the right 

side up) [Figure 41. 

b Biopsy punch made by Baker/Cummins, Miami, FL 

c Manghini liver biopsy needle, Kormed, Minneapolis, MN 
Liver biopsy in the California sea lion is performed on the right side of the 

body wall. To locate the exact site, place the right foreflipper back and flat against the 

right body wall. Where the foreflipper flexes at its carpus [Figure 41 is the insertion 

point. Be certain to position the right foreflipper mid-way between the dorsal spine 

and ventral body wall to avoid penetrating the pleural space (lung field). Insert the 

needle via a short, quick, penetrating action. Try not to penetrate the pleural space 

(lungfield). The biopsy core is aseptically transferred to a container of 10% formalin 

for fixation prior to histopathologic examination. 

I 

Figure 4 : Liver Biopsy Site ............................... ............................... 

( KEY needle insertion point 

BONE MARROW: 

Bone marrow biopsies can be used routinely and provide excellent diagnostic 

information regarding pinniped immunologic status, hematologic capacities and 

potential neoplastic (cancerous) activities. In pinnipeds, palpate the fkmoral 

trochanter. Over this area, make a 2 cm surgical incision. A Jamshidid bone 

marrow needle is used to penetrate the center of the trochanter area to a level of 

about 3 cm. Remove the bone marrow biopsy needle stilette and aspirate bone 

marrow material with a sterile 12 cc syringe. 

Place a small portion of bone marrow tissue on a clean microscopic slide. 

Smear it out (as with a blood smear), and allow to air dry. Take the remaining 

biopsy material and place in a vial of 10% formalin. 

d Jamshidi bone marrow biopsy needle, Kormed, Minneapolis, MN 
MISCELLANEOUS BIOPSIES: 

Fine-needle aspiration biopsies11 can be used to collect cytological materials 

from external lesions, or from any suspected internal areas. Aspiration biopsies are 

easy, quick, and capable of yielding excellent diagnostic information on the spot, 

assuming microscopic equipment is available. 

Use a sterile set-up of a 22- or 25-gauge needle attached to a 12 ml syringe. 

Insert the needle into the area or lesion you wish to investigate. Aspirate, rapidly 

pulling the plunger at least half way up the barrel, creating negative pressure. 

Without removing the needle, reposition it in another location within the area of 

consideration, and repeat. Gently withdraw the needle (still attached to the syringe) 

from the lesion. Gently pull back on the plunger, bringing I to 2 ml of air into the 

syringe. Expulse the material in the needle hub onto at least three clean glass 

microscope slides. Make a smear of the aspirate on each slide, as you would for 

blood count differentials. On one of the slides, place one or two drops of new 

methylene blue stain and a coverslip, or, as with a blood smear, stain the slide with 

Diff-Quik. Send the other two air-dried needle aspirate smears to the laboratory for 

further analysis. It is best to call the laboratory prior to obtaining the specimen to 

ask the preferred method of aspirate collection and preparation. 

LIFE HISTORY INFORMATION 

PERTINENT INFORMATION: 

Pinniped life history information, including age, sex and reproductive history 

deduced from ovarian structures, can be critical for assessing population trends, 

forensic data and contaminant analysis data. 

Determine the genus and species of the pinniped (See references under 

Pinniped Identification). 

Determine the sex of the pinniped. Differentiating male from female 

pinniped is most easily accomplished by rolling the animal over on its back and 

examining for a penile orifice just caudal to the umbilical scar. Adult male otariids 

may be identified by noting the scrotal sac posteriorly, but if the sac is drawn close to 

the body, this may be difficult; remember, that in phocids there is no scrotal sac. In 

young animals, it may be difficult to actually see the penile orifice through the 

lanugo coat; however, there is often urine staining, and the os penis (penile bone) 

can always be palpated through the skin [Figure 51. 
Weigh the pinniped, and record the weight on both the standard 

measurement and necropsy forms. As you perform a necropsy, specific organs 

should be weighed as well (See Standard Necropsy Fo.rm and Specimen Checklist for 

Histopathology on pages 21-23 & 52). 

Determine the of the pinniped, if possible. Age determina8ion without 

history is difficult and speculative. The best indicator of age is the tocrth. Remove a 

canine or postcanine tooth, and process it appropriately (see Section on 

Tooth/ Bone/Skeletal Preparation). 

Other specimens to collect that will assist the pinniped life histbrian in data 

collection arel2: 

(1) Whole skull (for morphometrics); 

(2) Lower jaw (with teeth present); 

(3) Stomach and stomach contents (for feeding habits, nutritional information 

and biotoxicant presence); 
(4) Reproductive organs (ovaries can indicate pregnancies, testes can 

indicate maturity); 

(5) Fetus (for developmental characteristics and deformities); 

(6) Milk (for nutritional status of mother and for handraising formulation); 

(7) Parasites (for developing baseline and evolutionary information, including 

nutritional status and feeding and haul-out habits); and 

(8) Prey items (intact species from the pinniped's habitat for comparison purposes, 

and for feeding habit information). 

[See Table 3 and Section on Specimen Collection] 

PINNIPED STANDARD MEASUREMENTS: 

To the clinician, technician or volunteer, the importance of taking standard 

external measurements on pinnipeds is not altogether clear. But without this 

information, life history scientists, policy makers, and conservation specialists, who 

must research and protect populations of animals in the wild, have greater difficult 

in devising management decisions. It is extremely important to these decisionmakers to know whether the animal being examined is normal size for its age, in 

good flesh, and of correct morphometric dimensions for its particulak- genus and 

species. 

Standard measurements in pinnipeds should be performed op live animals 

at the time of initial examination if possible. In the live animal, it may not be 

feasible to collect more than the standard length, curvilinear length, axillary girth, 

umbilical girth, flipper lengths and widths and tail length. Record all data in metric 

measurements. The numbers or letters next to the specific measurements refer to 

the numbers and letters in Figure 6. Always be certain to weigh the animal, sex the 

animal, and make an estimate of the animal's age. 

In a dead animal, be certain to record all life history information, as well as 

take a right upper canine tooth for exact aging of the animal. The importance of life 

history information cannot be overemphasized, as it characterizes whole 

populations of marine mammals, and allows the scientists to begin to develop a 

scenario as far as population dynamics are concerned. 

Record the number and type (incisors, canines, post-canines) of teeth in upper 

and lower arcades (jawbones) to aid in further pinniped life history identificationl3. 
16 

TABLE 1: PINNIPED STANDARD MEASUREMENTS 

(record in kg and cm) 

0- Body Weight 

1. Standard Length 

2. Curvilinear Length 

3. Axillary Length 

4. Umbilical Girth 

5. Eye Length ---------------- 

6. Ear Length ---------------- 

7. Anterior Length of the 

Foreflipper 

8. Axillary Flipper Length 

9. Widest Foreflipper Length 

10. Anterior Length of the 

Hindflipper - 

(record in cm) 

11. Widest Hindflipper Length 

12. Tail Length --------------- 

Skull Measurements 

A. Rostral Width 

B. Interorbital Width 

C. Zygomatic Width 

D. Cranial Width 

E. Condylobasal Length 

F. Rostral Length 

G. Length of Upper 

Postcanines - - 

Internal Measurement 

13. Blubber Thickness 
The following shows required standard body measurements for pinnipeds [Figure 61. 
STANDARD NECROPSY PROCEDURES IN PINNIPEDS 

GROSS NECROPSY 

EQUIPMENT NECESSARY: 

Camera, 35mm, with 50mm and macro (close-up) lenses, flash and extra 

batteries; Video camera, if available (the video camera should not 

substitute for clear and accurate photography) 

Protective clothing (plastic gloves (preferably talc-free), boots, aprons) 

Surgical mask and protective eyeglasses (to guard against public health risk) 

Record sheets (including standard necropsy form and standard 

measurement sheet), soft lead (no. 2) pencils and clipboard 

Walk-on scale and wheelbarrow for weighing whole animal 

Flexible and metal tape measures (minimum 5 yards, with centimeter markings) 

Plastic/wooden ruler, graded in centimeters and inches 

Tooth dental elevator 

Dissecting tray or board 

Sharp high quality stainless steel knives (curved and/or straight, 

minimum 6 inch blade) 

Sharpening steel 

Scalpel blades (assortment of #10s and #15s) and handle 

Small and large dissection scissors 

Long and short-handled, tissue and rat-toothed forceps 

Pruning shears, bolt cutters, or oscillating (Stryker) or electric saw (for 

cutting bone and cartilage) 

Bone rongeurs (for removing small bony fragments) 

Heavy duty rubber gloves (able to be disinfected and cleaned) 

Cotton string (for tying off fluid filled hollow organs, i.e stomach, etc.) 

Spring or platform weighing scale (20 kg capacity) 

Postal scale or other weighing device (in mg & gram weights) 

Paper towels 

Tyvek Labels and indelible ink pens/ permanent markers 

(non-smearing) [see example of label front & back below] 

Containers for collected samples (see Section on Shipping) 

Fixatives (10% Formalin; also see Appendix A) 

A1c:ohol (70% ethanol) 

Various sterile blood collection tubes, syringes and needles 

Whirl-Pake or zip-lok plastic bags 

Aluminum foil (heavy duty) 

Covered barrel for solid waste disposal 

c Whirl-Pak, Fisher Scientific, Englewood, CO 
Speclmen Label 

Date: -------------- From: ------------------- 

---a-a------------- 

Animal ID Number= --------- 

Genus & Species: ------ 

- 

Sex: ------ Age: -------- <over> 

(front) 

To: 

------ 

------------------- 

------------------- 

Comments ------------- ...................... 

...................... 

...................... 

Contact for Add'l Info 

(tell: ---------------- - 

(fax): ------------------- 

Information (bac Label k) 
STANDARD NECROPSY FORM: 

A great deal of important comparative data can be lost if the clinician or 

technician concludes that there is "not enough time" to write up a report, or "the 

findings are not significant enough to record. If there is enough time to perform a 

necropsy, writing the report is part and parcel of that necropsy. The importance of 

records, particularly in a court of law, cannot be overstated. 

The following standardized necropsy form is recommended for a methodical, 

step-by-step assessment of organs and lesions post-mortem. Tissue samples should 

be routinely collected from all organs, whether thev appear grosslv normal or 

abnormal. When sampling abnormal tissue, be certain to collect adjacent normal 

tissue. This standardized necropsy form allows a worthwhile, item by item 

examination that is relatively rapid and error-free; it also provides consistency, 

whether the reporter is clinician, technician, or volunteer. 

The person performing the necropsy must alwavs leaiblv sign the necropsv form, 

so that examination information can be traced back, if necessary, at a later date. 

Some facilities require that necropsy reports dictated and/or transcribed during 

actual necropsy procedures be formally typed up for distribution and/or reporting 

purposes. The original handwritten and signed document prepared at the time of 

necropsy should routinely be kept on file for future reference, be it of a legal or 

scientific nature. 
I . Administrative Information I. Date 

2. Facility Name 

3. Pinniped Genus 4. Species 5. Sex 

6. Animal Identification Number 7. Other Id (tags etc.) 

8. Necropsy Personnel 9. Attending DVM 

10. Time of Death (24 hr) 11. Died/ Euthanized /Other (explain) 

12. Time of Necropsy~4 hr) 13. Location of Necropsy (city,county,state) 

14. Photographs Taken 15. Name of Photographer 

16. Weight of Animal (kg/lb) 17. Age of Animal/Tooth Taken (circle one) 

I Ancillary Administrative Information 

Stranding Information: 

18. Date of Stranding 19. Time of Stranding (24 hr) 

20. Exact Location of Stranding 

21. All same species of Pinniped? 22. Number of Animals 

23. If not all same, numbers by Genus and Species 

Weather Information: 

24. Weather Conditions (circle as many as necessary) 

clear, partly cloudy, overcast, rainstorm, snowstorm, windstarm, 

thunder, lightning, sand storm, fog, other conditions (explain) 

25. Wind Direction 26. Wind Speed 

27. Tides 28. Swells/ Waves 29. Moon 

30. Water Temperature 31. Air Temperature 

Other Historical Information: 

32. Fish run/Plankton bloom ongoing? (explain) 

33. Any geographical abnormalities near stranding site? (explain) 

34. Any geological abnormalities near stranding site? (explain) 

35. Any industrial/ urban /agricultural enti ties near site? (explain) 

36. Any other causes of concern near the site? (explain) 

37. Any medical problems obvious in animals? (explain) 

38. Any clinical signs present in animals? (explain) 

1 39. Any other historical information not already included above? (explain) 
II. Gross Necropsy Information I 

General Condition (circle applicable descriptions) 

Carcass condition Code (2,3,4,5) 

Postmortem condition (good, fair, poor, mummified) 

Nutritional state (overweight, fat, thin, emaciated) 

State of hydration (good, slightly dehydrated, moderately dehydrated, 

severely dehydrated, unknown) 

Haircoat (normal, hair loss, oil, molting, scruffy, parasites, 

Skin (normal, cracking, bleeding, dried, moist, smooth, rough, wounds, 

infection, masses, parasites, scars) 

Nails (normal, present, absent, torn, bleeding, cracked, crushed) 

Peripheral lymph nodes (palpable, obvious, unnoticeable, other) 

Star (with an asterisk *) each tissue taken for analvsis I 

Body Orifices (describe what you see) 

Mouth and teeth 

Nose 

Eye, left 51. Eye, right 

Ear, left 53. Ear, right 

If male, Penile orifice 55. If female, Vulva 

Anus 

Abdominal Cavity (describe what you see) 

Relative position of organs 

First impressions on opening the abdominal cavity 

Describe Conditions of Organs and Tissues, Any Lesions 

Peritoneum 

Stomach 

Duodenum 

Small intestine 

Large intestine 

Mesenteric lymph nodes 

Pancreas 

Spleen 

Liver 

Gall bladder and bile duct 

Adrenal gland, left 70. Adrenal gland, right 

Kidney, left 72. Kidney, right 

Ureter, left 74. Ureter, right 

75. Bladder and urethra 
Reproductive Organs 

FEMALE-- MALE-- 

76. Ovary and oviduct, left 82. Testicle and vas, left 

77. Ovary and oviduct, right 83. Testicle and vas, right 

78. Uterus and cervix, left 84. Prostate 

79. Uterus and cervix, right 85. Penis and os penis 

80. Vagina and vestibule . 86. Prepuce 

81. Mammary glands 

Thoracic Cavity (describe what you see) 

87. Diaphragm 

88. Relative position of organs on opening the thorax 

89. First impressions on opening the thorax 

Describe Conditions of Organs and Tissues, Any Lesions 

90. Lung, left 94. Mediastinal lymph nodes 

91. Lung, right 95. Pleura 

92. Ribs, left side 96. Thymus 

93. Ribs, right side 97. Brown fat 

The Neck (describe what you see) 

98. T1.1yroid and Parathyroid, left 

99. Thyroid and Parathyroid, right 

100. Esophagus 

The Pluck (describe what you see) 

101. Tongue 

102. Laryngeal Area 

103. Trachea 105. Mainstem bronchus, right 

104. Mainstem bronchus, left 106. Bronchioles 

The Heart (describe what you see) 

Pericardium 

Myocardium 

Endocardium 

The Valves: The Vessels: 

Tricuspid 114. Pulmonary vein 

Pulmonic 115. Pulmonary artery, right 

Mitral 116. Pulmonary, left 

Aortic 117. Dorsal aorta 

The Chambers: 

Right atrium 119. Left atrium 

Right ventricle 121. Left ventricle 

The Nervous System (describe what you see) 

122. Brain 

123. Spinal cord 

124. Peripheral nerves 
IN GENERAL: 

Be certain when performing any necropsy or tissue collection procedures on 

pinnipeds to wear protective gear, including surgical gloves, mask, and apron, if 

possible. Thoroughly disinfect instruments before and after the procedure. 

Protective gear and disinfection are recommended to minimize the potential risk of 

necropsy personnel acquiring any microbiological or chemical contaminant from 

the pinniped being exarnined.14 

Gross pathological protocol is an important first step for the diagnostician in 

determining potential causes of death and disease. Necropsy (from the Greek "to 

view death") or post-mortem examination involves a complete external and 

internal examination of the dead animal. 

Gross necropsy, as with other forms of medicine and forensics, is as much an art 

as a science. Before any description can be written, a thorough and consistent 

examination of all organs and lesions (called a necropsy, or animal autopsy) must be 

performed. An assistant can be most helpful; the assistant can write and weigh 

organs, as you perform and describe the lesions and organs you see. Whether you 

have an assistant or not, it is most important to write the report as you perform the 

necropsy or directly afterward, so as not to forget or omit any inform~tion from the 

description. Descriptions that are the most useful are simple, clear, and without 

interpretation. Describe only what you see. Perform the necropsy in a methodical 

manner, one system at a time, in the same order each time. Always describe each 

organ or lesion's size, color, shape, consistency, location, texture on cut surface, odor 

(if any), general description and weight. Record size and weight metdically, to 

coincide with observations others might make on similar animals. 1f you see no 

obvious abnormalities grossly, record "no significant lesions" (NSL) or "no 

significant observations" (NSO). As the examiner performs more and more 

necropsies, the powers of observation will improve and become accurate, precise 

and succinct. Practice makes perfect. 

GENERAL EXTERNAL EXAMINATION: 

Examine the skin, appendages (flippers, tail, head) and body orifices for wounds, 

oil/tar, hair loss, external parasites, texture changes (cracking, bleeding, drying, 

moisture, smoothness) and condition of the nails (present, absent, torn, bleeding). 

Unless the species of pinniped you identify is listed as threatened or endangered, 

remove a canine tooth for aging purposes. If the exact age of the pinxtiped is known, 

you need not remove a tooth. If the pinniped is a listed species, do not remove a 

tooth until you contact your regional NMFS office for further instructions. 
In general, for ease, efficiency, and conciseness, the person performing the 

necropsy examination should examine the following decision matrix to determine 

tissues and collection media that might be necessary to help the clinician and/or 

diagnostician determine the cause or causes of death in the pinniped under 

consideration [Table 31. 

At the time of gross necropsy, at the time each lesion, organ or tissue is examined-- 

1. Collect sample for histopathology, including normal and abnormal appearing 

tissue in the sample (see pages 51-52); 

2. Collect representative parasites seen grossly (see pages 53-54); 

3. Collect, aseptically (without contamination), sample for microbiology, including 

bacteriology, virology and mycology (see pages 54-59); 

4. Collect appropriate tissues or specimens for toxicological analysis, using clean 

cutting instruments, with all instruments being wiped clean between 

samplingf (see pages 59-63); 

5. Collect any other tissues or specimens that might be necessary for the 

clinician/diagnostician to make a diagnosis (see pages 64-65). 

Each of these 5 procedural areas will be discussed independently and in detail 

beginning on page 51 below, but remember to consider them as a continuum if the 

cause of death is unknown. Collecting duplicate samples, sending one set of tissues 

for analysis, and archiving the second set on-site, insures the availability of samples 

at a later date if additional analysis or rechecking of results is necessary. These rules 

are especially important if there is any concern that the data from tissues or 

specimens collected may be necessary for use in a court of law. 

f For optimum results use high carbon stainless steel instruments cleaned 

with deionized water 
Table 3: Decision Matrix for Pinniped Tissue and Specimen Collection 

USE THE 

Fixatives 

Containers 

Other 

Temperature 

X 

X 

X 

X 

X 

XX 

X 

FOLLOWING PREPARATION: 

10% Formalin 

70% Ethanol 

2.5% Potassium Dichromate 

AFA Solution 

Physiologic Saline 

3% Glutaraldehyde 

Trisodium Phosphate 

Covered Plastic Jar 

Sterile Swab/Culturette 

Sterile Red Top Tube 

Sterile Purple Top (EDTA) 

Dermatophyte Test Media 

Michel's Media 

Screwtop Polyethylene Vial 

Acid-rinsed Glass Jar 

W hid-pak/Zip-loc Bags 

Aluminum Foil 

Blue Top Tube (citrate) 

Whole Blood 

Serum 

Plasma 

Biopsy 

Tissue 

Room 

X 

X 

X 

Refrigerated (4 C) 

Frozen (-20 C) 

Dry Ice - 

X 

X 

X 

X 

XXX 

XXXXXX 

XXXXXX 

X 

XX 

XX 

X 

XX 

X 

. 

XXX 

X 

X 

X 

X 

X 

X 

X 

X 

X 

X 

X 

X 

XX. 

X 

X 

XXXX 

XXXX 

-- 

I 

X I 

- 

1- ----I 

X 

X 

X 

X 

-, 

--- X 

X 

X 

X 

IX 

X 

X 

XX 

X 

X 

X 

X 

X 

1 

X 

X 

I 
DESCRIPTORS FOR USE DURING NECROPSY REPORTING 15 

DESCRIBING ANATOMICAL LOCATIONS: 

Initially describe each organ or lesion by its location within the body cavity, as 

well as its relationship to other organs or structures. Become familiar with 

anatomical descriptors for organ and organism position [Figure 71: 
Dorsal 

Ventral 

Cranial 

Caudal 

Lateral 

Medial 

Proximal 

Distal 

Anterior 

Posterior 

- of, toward, on, in or near the back 

- pertaining to the lower side or undersurface 

- toward the head 

- toward the tail 

- of, relating to, or situated on the side(s) 

- relating to, lying or extending toward the middle 

- nearer to the main body of the organ or organism 

- away from the main body of the organ or organism 

- on the forward leading edge 

- on the backward trailing edge 

DESCRIBING SIZE: 

Always accurately describe size. Do not say "small", or "larger than normal", 

record metric length, width or diameter, and metric wet weight (this is the weight 

when the organ is freshly removed). A millimeter/centimeter ruler should be on 

hand for accurate measurement. If a metric ruler is not available, score your 

forefinger from joint to joint, so that you can estimate centimeters. 

DESCRIBING COLOR. 

The easiest way to describe color if it is not a standard primary color, is to use a 

descriptor that refers to a common everyday item (food is a good choice). In 

addition, other adjectives and adverbs, such as mottled, shiny, pale, sweaked, 

blotchy, and dark are words used to describe organ or lesion color. 

DESCIUB:NG SHAPE AND FORM: 

Describe what you see, using descriptors. The following words13 are merely 

suggestions: oval, ovoid, spherical, round, egg-shaped, discoid, square, cuboidal, 

rectanguliw, triangular, wedge-shaped, spindle-shaped, long, slender, flattened, 

domed, swollen, bulbous, indented, lobulated, lace-like, spongy, narrowed, patent, 

dilated, open, tortuous or branching. Cut-surfaces can be described as swollen, 

bulging, shiny, dull, eroded, glistening, scaly, pitted or oozing. The terms 

amorphous, shapeless or irregular can be used if there are no definite shapes or 

patterns, or the shape is too erratic to describe. If you have performed a number of 

necropsies and recognize that the shape of a particular organ is normal, you may 

record a phrase such as "of the usual shape". 
DESCRIBING CONSISTENCY: 

This is an easy category. Descriptions here may be plain and simple, such as 

firm, hard, soft, flabby, fluid-filled, sharp-edged, or friable (falls apart with any 

amount of pressure). If any fluids are present, they should also be described, such as 

cloudy, turbid, clear, thick, bloody, dark or pus-like. 

DESCRIBING ODOR: 

Do not overlook your sense of smell; it can be an excellent indicator of 

pathological processes. With a fresh specimen, there is rarely any obvious odor. 

Sweet, sour, rancid, ammonia-like, fetid, putrid, fermentative, petroleum-like, or 

similar descriptors are useful. 

EXTERNAL EXAMINATION OF THE CARCASS 

First 

Weigh the animal and take standard pinniped measurements. 

Note the presence of any tag/identifying marks on the pinniped. 

Collect apropriate samples [See Table 3 and Section on Collection, Preparation 

and Preservation of Tissues and Specimens] . Mark Samples taken on checklist 

for Histopathology. 

Note the condition of the carcass (Code 2,3 or 4). Note the anim,al's general 

body condition; is the animal in good flesh, thin, or emaciated (ribs showing)? Is 

the animal dehydrated? The skin turgor/fallback test can be used to check 

hydration, but if the condition of the carcass is not absolutely fresh, or there is a 

large amount of blubber present, this test may be unreliable. If performed, a flap of 

skin between the animal's shoulder blades is lifted and left to fall back down; rapid 

fall indicates good hydration; if the flap hesitates before falling back flush with the 

body, the animal can be considered dehydrated. 

Hair Coat 

All pinnipeds, except walruses, have hair overlying their skin. Odobenids 

(walrus) are hairless with raised, irregular "blebs of scar tissue-like" skin. Most 

otariids and phocids have .dense hair coats of varying lengths and densities, 

depending on the particular species. Phocids and otariids, adults and neonates, go 

through hair coat molts, with the phocids more likely sloughing hair in sheets, 

while otariids can shed and regrow a portion of their hair coats almost 

unperceptively each year; note any hair coat changes. 
Do not be confused by droplets of water on pinniped pelage; these areas can 

appear dark and shiny, but should not be confused with actual skin or hair coat 

lesions. 

Body Orifices 

All seven body orifices (mouth, nose, eyes, ears, penile orifice, vulva, anus) 

should be examined for wounds, parasites, pus, blood or other abnormalities. 

In the oral cavity, examine the tongue, the teeth (which can be stained a blotchy 

black or brown in pinnipeds), and note any abnormalities. If the animal is very 

freshly dead, note the color of the mucous membranes (pink, white, blue, red). 

The nose in elephant seals often has a normal white, pasty, nasal discharge. 

Other phocids, otariids, or odobenids do not normally have a nasal discharge. Look 

closely fbr parasites, discharges, wounds, foreign bodies, or other lesions in the nasal 

passages. 

Pinniped eyes are not easily opened. The pinniped eye has a third eyelid located 

medially and ventrally. In each eye, note any discharge, wounds, abrasions, foreign 

bodies, or other lesions. 

The ear in phocids and odobenids is no more than an external hole on each side 

of the head. In otariids, each ear has an external flap, called a pinna, which differs in 

length and shape depending on the species. For each ear, note any discharge, 

wounds, abrasions, foreign bodies, or other lesions. 

In the male pinniped, posterior to the umbilicus, is the penile orifice, where 

urine and sperm exit the body. The penis is hidden under subcutan~us fat, rather 

than hanging ventrally as in terrestrial mammals. Note any dischar$e or swelling 

at the orifice. Extend the penis by grasping the os penis. Note any lesions. 

In th.e female, just ventral to the anus, is the vulva. Note its color, shape, and 

any swelling, discharge, wounds or other lesions that might be present. 

The anus is located under the tail of the pinniped; it is the most dorsal orifice 

located on the animal's posterior. Note the presence of any discharge, diarrhea, 

wounds, or other lesions. 
Peripheral Lymph Nodes 

Peripheral lymph nodes are not easily located in pinnipeds due to either 

subcutaneous fat or to thick pelage (hair coat). If obvious on external examination, 

be sure to note lymph node location and any abnormalities. 

Mammary Glands 

Female pinniped mammary glands lie in the subcutaneous fat under the skin 

and tend to follow the general contour of the abdominal body wall. They are not 

normally visible unless there is milk, infection or other abnormalities present. 

Normally, the nipples are retracted under the hair coat and are often described as 

"inverted. Otariids (seal lions, fur seals) and odobenids (walrus) have four 

mammary nipples (teats), two anterior to the umbilicus and two posterior to the 

umbilicus, one on each side of the ventral midline. Phocids (true seals) have two 

abdominal mammary teats, either two directly adjacent to or two directly posterior 

to the umbilicus. 

Scrotum 

Testicles sit in a scrotal sac only in the otariids, and because the Sac is closely 

adhered to the body wall, is not easily visible. Phocid testicles are not present in a 

scrotal sac but lie deep in the subcutaneous fat of each inguinal area. 

INTERNAL EXAMINATION OF THE CARCASS 

Collect appropriate samples [See Table 3 and Section on Collection, Preparation 

and Preservation of Tissues and Specimens] .. Mark Samples taken on checklist for 

Histopathology as appropriate. 

Positioning the Carcass 

The pinniped is positioned in dorsal recumbency (the animal is on its back and 

symmetrically positioned with it four appendages out to the sides) [Figure 81. 

OPENING THE ABDOMINAL CAVITY 

Using a knife (if the pinniped is large) or a #10 scalpel blade (if the pinniped is 

small) incise (cut) the skin on the ventral midline just cranial to the umbilicus. 

Continue to incise cranially along the ventral midline to just under the level of the 

chin [Figure 8 63 I]. As you make the incision, extend it into the blubber layer of the 

animal if it exists. -- NOTE: Thin or emaciated pinnipeds may not have a thick 

blubber layer; also fur seals will not normally have a blubber layer. In these cases, 
take care not to incise the chest or abdominal cavities as you make your skin 

incision. -- Extend the skin incision similarly, but caudally from the umbilicus 

[Figure 8 @ 21. -- NOTE: If the animal is a male, avoid the os penis and penile orifice 

by taking the incision to the left around these organs. -- Continue the incision 

caudally to the left inguinal (groin) area [Figure 8 @ 31. 

Notice where the thorax stops and the abdomen begins in a pinniped. Palpate 

the level at which the ribs stop, approximately two-thirds of the way between the 

head and tail. Just caudal to the last rib is the start of the abdominal cavity. 

Next, make skin flaps on either side of the ventral midline incision, by incising 

perpendicularly to the midline, both from the chin area to the axillae (armpits) on 

either side, extending these cuts toward the dorsum [Figure 8 @ 41, and also from the 

inguinal areas on either side toward the dorsum. Record the color of the blubber 

and measure the thickness of the blubber layer (the blubber layer is the fatty, adipose 

tissue layer usually attached directly to the skin as you pull back the flaps). Using a 

metric tape measure, determine and record the depth of the blubber at its thickest 

point along the ventral midline incision. You may need to gently dissect downward 

through the blubber, avoiding penetration into the abdomen or thorax, by using 

blunt-tipped scissors [Figure 91. 

Once the blubber measurement is taken, record it on your Pinniped Standard 

Measurement form (page 16), then pull back the skidblubber flaps to expose the 

thoracic and abdominal walls. 
Underneath the blubber is a tough, sinewy tissue. The tissue caudal to the 

ribcage is the muscle overlying the abdominal peritoneum. The peritoneum lines 

the abdominal cavity, and is in two layers, the outer (ventral) peritoneum (usually 

covered by a thick layer of muscle) and the inner (dorsal) peritoneum (often only 

noticed firmly adhered to the ventral peritoneum on reflecting back the peritoneal 

layer with rat-tipped forceps). 

Make a small, sharp stab incision with the scalpel blade through the muscle and 

peritoneum into the abdominal cavity. Place two fingers inside the hole, spreading 

the edges laterally (out to each side) and tenting the tissue up. Using blunt-tipped 

scissors, incise caudally, cutting through both muscle and peritoneum. 

Within the abdominal cavity, note the relative position of all organs. Record 

size, color, shape, consistency of each organ one by one as you systematically 

examine the abdominal contents. 

GENEIUL CONDITIONS AND OBSERVATIONS ON OPENING THE ABDOMEN: 

The abdominal cavity of a pinniped, is a small compartment, relative to 

terrestrial mammals, occupying only about the caudal 1 /3 of the animal's body 

length. Overlying the majority of organs in the cranial abdomen is 8 strong, lacy, 

somewhat transparent tissue called the omentum. The inner (dorsal) peritoneum 

(abdominal lining) is very strong and very vascular, with, especially in the larger 

pinnipeds, a veritable net of blood vessels (called the rete mirabilae, translated as 

marvelous net) coursing along the inside lining of the dorsal peritoneum. 

Peruse the relative position of each organ you can see, noting any abnormal 

relationships, such as displacement, twisted or entrapped intestinal loops or organ 

size. Note presence, quantity and/or quality of any fluid within the abdominal 

cavity. 

Also note the general condition of the internal organs and tissues. In general, 

this observation of general condition should correspond with the external condition 

of the carcass (Code 2,3,4 or 5), which you recorded at the start of the necropsy 

procedure. Note the relative degree of tissue decay or decompositi~n (autolysis). If 

you observe other conditions, such as generalized hemorrhage (bleeding into organ 

walls or other tissues), generalized icterus (yellow jaundice) or generalized paleness 

or pallor to the tissues, note them at this time. 

STOMACH: 

In pinnipeds, unlike cetaceans, the stomach is a single-chambered organ. On 

incising the stomach wall, there are large, often reddish, rugal folds. Here and in all 

subsequent section of the gastrointestinal tract, be certain to note any obvious 
ulcerated or bruised areas, any hemorrhage, any possible foreign bodies, or any 

parasites that might be present (some internal parasites may be minute, so look 

carefully; a hand lens or magnifying glass can be helpful). 

If information on food consumption patterns is required, collect fish otoliths 

(ear bones, specific for each fish species) for identification by life historians. If whole 

intact prey (food) items are present in the stomach, collect these in sterile zip-loc 

bags and freeze for identification and/or contaminant or biotoxin analysis 

(see pages 59-63). 

In a normal pinniped, you will note that the stomach lacks an obvious pyloric 

region (in terrestrial animals the pylorus is a glandular area found at the point 

where food materials exit the stomach and flow into the small intestine). 

SMALL INTESTINE: 

The duodenum makes up the initial portion of the small intestine. The 

duodenum surrounds the pancreas (see below) as it courses caudally toward the 

jejunum and ileum (the successive portions of the small intestine). The pinniped 

small intestine in its entirety is not only much longer but more coiled than that 

seen in terrestrial mammals. In an adult California sea lion, small intestinal 

lengths up to 33 meters have been recorded. 

LARGE INTESTINE: 

In pinnipeds there is no true cecum, as is seen in most terrestrial mammals; 

instead, in pinnipeds, there is simply a small diverticulum (or sac-like pouch) 

which denotes the start of the large intestine. The large intestine (or colon) is 

exceptionally short in pinnipeds, usually less than 35 cm in length depending upon 

the age and species of the pinniped. 

MESENTERIC LYMPH NODES: 

The mesenteric lymph nodes are located centrally within the mesentery (the 

stained-glass appearing connective tissue that holds all the intestinal loops in 

alignment). Unlike terrestrial mammals, these lymph nodes are very large (8 to 9 

cm in length) and oblong in shape; they are also very "meaty-appearing' and 

usually bunched up together in the central portion of the mesentery, rather than 

spread throughout. A thick capsule surrounds each lymph node; check to see how 

closely adhered the capsule is to the lymph node by slitting the capsule and 

attempting to peel it back. Slice through a representative sample of lymph nodes, 

noting any hemorrhage, bruising, exudate or bulging on cut surface. 
PANC.REAS AND PANCREATIC DUCTS: 

Gently pick up the duodenum (the beginning of the small intedtine) in your 

fingers, and follow it along noting the pink meaty tissue lying adjacent to the 

duodenum. This is the pancreas; in the pinniped, it is a very large organ located 

"in the embrace of the duodenum". Visually examine and manually palpate the 

pancreas, noting any apparent nodules, masses, hemorrhages or exudates. 

Note the main pancreatic duct (the tube running centrally through the soft 

tissue mass of the pancreas) and the several lesser ducts (branching off from the 

main). Once visually examined, make a 6 cm long cut in the lateral duodenal wall. 

Gently milk the main duct of the pancreas with your right hand as you use your left 

hand to somewhat elevate the duodenum up for viewing. Pancreatic juices 

(generally white) should flow out the pancreatic duct and exit into the duodenum. 

Note duct patency (do the pancreatic secretions flow easily?) and the consistency of 

the fluid as it enters the duodenum. Then, using scissors, cut the main pancreatic 

duct open longitudinally, following its flow toward the duodenum, checking as you 

do so for any areas of hemorrhage, bruising, secretions, excretions, parasites or 

other abnormalities. 

SPLEEN: 

The pinniped's spleen is located, as in other mammals, on the right side of the 

abdomen (the left side as you look into the abdomen from the venltral midline). 

The spleen in pinnipeds, relative to domestic animals, is only of moderate size, and 

often has small, dark, homogeneously pigmented areas throughout, which contain 

hemosiderin (a non-pathologic iron pigment). Note any discolorations, raised 

areas, or apparent nodules or masses. Also note the thick, capsule covering the 

spleen; slit the capsule and attempt to peel it back. 

LIVER: 

The liver of the pinniped takes up a considerable portion of the cranial 

abdomen, lying against the muscular diaphragm and adjacent and dorsal to the 

stomach (deep in the abdominal cavity under the stomach at necrapsy). The 

pinniped liver is very large, multi-lobed, and has a considerable number of natural 

fissures (or slits) in it. To thoroughly examine the liver, make representative crosscutting slices ("bread-loafing") through all lobes, noting the organ's consistency as 

you do so. Also note any areas of discoloration, nodular occurrences, exudates or 

parasites within the parenchyma (deep tissue) of the organ,. 
GALL BLADDER AND BILE DUCT: 

The gall bladder is a green-tinted organ, usually located centrally or just offcenter, within the mass of the liver. Be certain to note the size and dative fullness 

of this fluid-filled organ. 

To check bile duct patency, use the 6 cm long incision made in the wall of the 

duodenum during examination of the pancreatic duct. Gently squeeze the gall 

bladder; bile (green fluid) should flow down the bile duct (tube) and exit into the 

duodenum in the same area as the pancreatic duct entered. Note bile duct patency 

(does the bile flow easily?) and fluid consistency as it enters the duodenum. 

ADRENAL GLANDS: 

In the pinniped the adrenal glands are very large and oblong. Thqy are located 

adjacent to the cranial end (pole) of each kidney, but are much more ~bvious than 

in most terrestrial mammals because of their size. The adrenal gland$ are very 

meaty and, on cut surface, they bulge and exhibit soft, sometimes vacpolated 

centers. In addition, on normal adrenal gland cut surface, there is a lighter-colored 

cortical area surrounding an inner darker-colored medullary area. The right 

adrenal gland is generally larger than the left. 

KIDNEYS: 

Pinniped kidneys look similar to cow kidneys, in that they are very lobular 

(multiple 2 to 3 crn clumps of kidney all connected together). Both kidneys are 

located deep against the dorsal surface of the abdominal cavity, with the right 

kidney located slightly more caudally, relative to the left kidney. 

Each kidney is encased in an extensive venous net (called a superficial venous 

plexus); these venous nets tend to be more obvious surrounding pho~id (the true 

seals) kidneys rather than otariid (sea lions and fur seals) kidneys. Betause each 

lobule is composed of up to 100 individual kidney units (nephrons/rqniculi), on 

kidney cut surface each unit of the kidney exhibits a complete cortex (khe light 

colored tissue, located peripherally in each unit) and medulla (the dadk colored 

tissue located centrally in each unit). Slice each kidney longitudinally (from pole to 

pole), noting its consistency (excess fibrous tissue? mushy texture?), ar)y areas of 

discoloration, bruising, hemorrhage, exudate (pus-like material), or ~hether it 

bulges on cut section. Examine the pelvis (where all the collecting tubules and 

blood vessels exit the kidney) for the presence of calcified kidney stones, parasites, 

tumors, cysts or other abnormalities. 
URETERS: 

In the pinniped, each ureter (the urine draining tube) exits the kidney medially 

and very near the caudal pole of the kidney (not medially and cenQally as in 

terrestrial mammals). The left ureter is longer than the right ureter, simply because 

the distance from the left kidney to the bladder is greater than on the right. Palpate 

the entire distance of each ureter, from kidney to bladder, checking for patency or 

abnormalities such as pus and calcified stones, and take representative specimen 

samples for histopathology. Cut the ureters longitudinally, following them to, but 

not into the bladder. 

BLADDER AND URETHRA: 

In the pinniped, the bladder is a very small, highly muscular organ. It rarely 

will contain urine at post-mortem. If the bladder does appear to have urine in it, 

collect a sample of the urine using a sterile needle and syringe, and place the sample 

in a sterile red top tube for transport to a laboratory. 

Once a urine sample, if present, has been collected, cut open the urinary bladder 

and note the presence of any stones, blood, pus, or bruising, hemodrhages, growths, 

nodules or diverticulae (outpouchings) in the bladder wall. Check to be certain that 

the two ureters enter the bladder symmetrically on either side of the bladder neck. 

The urethra should exit centrally and caudally from the bladder to course through 

to the vagina in the female or the prostate and penis in the male. Check the urethra 

for patency and any abnormalities such as strictures, tumors, or urpthral stones. 

REPRODUCTIVE ORGANS: 

-- FEMALE 

OVARIES AND OVIDUCTS: 

Each of the two pinniped ovaries lie in a large, flattened pocket! (or bursa) at the 

start of each oviduct. In the neonatal female pinniped, the ovaries are large, meaty 

and close to adult size, but without any obvious structural elements. In adult 

female pinnipeds, examine both the left and right ovaries and oviducts. Describe 

any structures present on either ovary, recording the exact numbeq of each structure 

on both the right and left ovaries. Any fluid-filled, semi-transparknt "cyst-like" 

structures may suggest to the clinician that the female pinniped w8s in a follicular 

phase of her reproductive (estrous) cycle. In addition, if there are hard, circular, 

whitish to yellowish "nodules", or white, somewhat circularly scafred areas, these 

might suggest how long ago and how many young that particular female has 

produced. Note the number, size, color, and consistency of each structure on each 

ovary. 
Preservation of ovaries differs depending on the analyses to be performed. 

Histopathologic and life history ovarian specimens are removed intact and placed 

in a container of 10% formalin (see Appendix A). If ovaries are to be used for 

toxicologic analysis, they are removed, placed in a zip-loc bag and frozen at -20°C. Be 

certain to label the ovaries as left and right, and label the specimen plackage. 

Each oviduct4 bursa lies near the start of each uterine horn. Trace each oviduct 

at its origin, where the ovary is, caudally to the uterus. Examine the bursae and 

report any swellings or constrictions. 

UTERUS AND CERVIX: 

In the normal neonatal (less than 7 days of age) female pinniped, the uterus is 

hypertrophic (large and fleshy). In all female pinnipeds, the uterus i(s bicornate (i.e. 

it has two large horns, rather than a large body, as in humans), and f/he two horns 

along almost their entire length are separated by a well-developed sdptun~ (sheet of 

tissue). Phocids have a long urogenital canal with a prominent fold of tissue (called 

the hymenal fold) at its most caudal aspect. 

The cervix (where the uterus exits into the vagina) is a single canal in phocids. 

In otariids the cervix is double, that is, it has two channels that exit into the vagina. 

In the adult female pinniped, be certain to cut open the uterus and cervix and 

note any foreign materials, such as fetuses, pus (infectious thick odiderous material), 

serum or blood, as well as any fibrous tissue, strictures, masses or other mechanical 

obstructions. 

VAGINA 'AND VESTIBULE: 

In the female pinniped, the vagina widens somewhat from whe~te the cervix 

ends caudally toward the vestibule (the area just inside the external vulva). 

Examine the vagina and vestibule for tonicity, color, consistency and the presence of 

any growth or nodular structures. 

-- MALE 

TESTES AND VAS DEFERENS: 

Externally, only testes of male otariids (sea lions and fur seals) ca~ be found in a 

scrota1 sac, and even in otariids this sac is often closely adhered to thei body wall. All 

male phocid (true seals) and odobenid (walrus) testes are located in dhe inguinal 

area (groin), outside the abdomen, but deep under the skin and blubber, so as not to 

be visible externally. 
Both the left and right testicles of the male pinniped should be examined. Note 

the relative abdominal or inguinal position, and measure the length and width of 

each testicle. Describe any structures on or under the surface capsule of each testis. 

Note blotchy, scarred, hemorrhagic, or other non-uniform areas if present. 

Follow the vas deferens from each testicle down to the level of the prostate. 

PROSTATE: 

The prostate is not an easy organ to visualize in the male pinniped. It is often 

not only diffuse, but buried deeply and dorsally in the abdomen, just ventral to the 

pelvic bones and colon. It may be easiest to locate by following the short colon 

through the pelvic inlet. If this is impossible to do manually, the animal may have 

to be turned over (on to its ventral surface) and some of the pelvic bone removed 

dorsally to visualize the organ. 

PENIS: 

In the pinniped, the majority of the penis (except for the penile orifice at the 

skin surface) lies underneath the skin and firmly adhered to the ventral body 

surface. The penis is composed of a single, centrally located, blooddfilled, spongelike tissue, called the corpus cavernosum, which forms the bulk of the penis. The 

male pinniped, whether phocid or otariid, has an os penis (a long, thin triangular 

shaped bone) located centrally and longitudinally in the penis, alosgside the urethra 

(which courses cranially at this point toward the urethral (penile) orifice); the os 

penis is situated halfway between where the penis originates at the ventral body 

wall and the penile orifice, where the urethra exits the body. Note any abrasions, 

lacerations, bruises, bleeding, scar tissue, growths or other penile dbnormalities. 

THE THORACIC CAVITY 

The thoracic cavity in pinnipeds, when compared to terrestrial mammals, is 

very large. In general, the pinniped thoracic cavity extends caudally incorporating 

approximately 2/3rds of the animal's total body length. Presumabqy, the thoracic 

cavity has not yet been invaded during the necropsy procedure to tihis point. On 

opening the thorax, describe the relative positions of the organs, as well as any 

abnormalities or the presence and quantity of free fluid (blood, serum, pus) in the 

chest cavity. 
DIAPHRAGM: 

Examine the diaphragm from the cranial abdomen. Note that the diaphragm is 

an immense muscular structure that separates the thoracic cavity from the 

abdominal cavity. From the abdominal cavity, using a scalpel blade make 

approximately a 20 cm incision in the diaphragm, while observing the lungs. As 

the thoracic cavity is entered, the lungs (if they are healthy) should collapse, as the 

positive pressure from ambient air invades the air-proof chest cavity. 

LUNGS AND BRONCHIOLAR LYMPH NODES: 

(See under "The Pluck" on page 43) 

RIBS: 

The ribs are entirely cartilaginous and can be cut with sharp scisgors or scalpel 

blade. If the ribcage needs to be opened, it may be easier in larger pihipeds, such as 

male sea lions, to use pin cutters (used in orthopedic surgeries) or heavy-duty 

garden shears to cut the ribs, even though they are cartilaginous. Cut the ribs close 

to their dorsal attachments to the spine and remove the sternum and ventral rib 

cage. 

THE NECK 

THYROIDS AND PARATHYROIDS: 

The thyroids and parathyroids are located as paired struchrres adjacent to the 

ventral surface of the neck, just caudal to the laryngeal apparatus. be thyroid 

glands are often pinkish in color and plump in size and consistency. Parathyroids 

are smaller and firmer, with a slightly darker color. Make cross sectional slices 

through both the thyroid and parathyroid glands, noting consistencyk texture, any 

discoloration, hemorrhage, nodules or exudate. 

ESOPHAGUS: 

As in terrestrial mammals, the esophagus runs longitudinally d~wn the left side 

of the pinniped's neck. However, in the pinniped, the esophagus is Very long and 

extremely distensible, in order to accommodate large fish that the adimal may 

consume. Once the pluck is removed, longitudinally incise represestative portions 

of the esophagus, and note any lesions or unusual occurrences. 
MEDIASTINAL LYMPH NODES: 

Mediastinal lymph nodes are located along the central midline in the cranial 

portion of the thoracic cavity. In pinnipeds, these lymph nodes, like those in the 

abdomen, are large, long, red and meaty. Cut representative cross sectional slices 

through these lymph nodes, noting any abnormalities. 

THYMUS AND BROWN FAT: 

The thymus and brown fat are present only in young pinnipeds. The thymus is 

a red and meaty structure found just inside the cranial thoracic cavity; the thymus 

lies over the heart just to the left of center midline. Note the size of the thymus. 

Brown fat is a dark yellow, thick, fatty tissue found in sheets subcutaneously 

over the back and the thoracic walls of the neonatal pinniped. Note the estimated 

quantity and location of the neonatal pinniped's brown fat. 

Slice through both the thymus and brown fat, noting consistencies and any 

abnormalities. 

THE PLUCK 

The organs of the head and neck along with the lungs and heart can all be 

removed together. 'The pluck", as this is called, is removed by peeling the skin off 

the ventral chin and neck. The muscles at the sides of the mouth are incised and 

the incision on each side is extended caudally to the level of the thgrax, and the 

already exposed cut ribs. The tongue is grasped and pulled caudally; as this is done 

the connective tissue and muscle still attaching it to the sides of the carcass's neck 

are cut. The entire tongue, larynx, trachea, esophagus, bronchi, luqgs and heart 

should come out as a unit for closer examination. 

TRACHEA AND MAINSTEM BRONCHI: [Figure 101 
The trachea bifurcates cranially into two mainstem bronchi. In otariids, the 

trachea bifurcates more cranially than in phocids. The right bronchus branches off 

about 2 cm cranial to the left bronchus. Both bronchi in their entirety are very wide 

in diameter and short in length. In all pinnipeds, except the walrusl, the trachea, 

bronchi and large bronchioles all contain cartilaginous rings; these $ings are 

incomplete and C-shaped, except in the walrus, where the cartilagiqous rings are 

complete. In otariids, but not in phocids, the cartilaginous rings even extend down 

the bronchioles into the terminal airways (the alveoli). Using scissors, cut 

longitudinally down the trachea and each of the mainstem bronchi, noting any 

foreign bodies, bruising, hemorrhage, parasites (such as lungworms), exudate, 

tumors or other abnormalities. 

-. 

LUNGS AND BRONCHIOLAR LYMPH NODES: 

ht bronchus 

The lungs in pinnipeds occupy the majority of the thoracic cavity. On opening 

the chest cavity, the lungs should appear pink in color. On cut surfbce, there should 

be air in the bronchioles and bronchi. Healthy lung tissue placed in water or 

formalin, should float. Make multiple cuts in the lung tissue, looking for any 

parasites, mucus production, pus, blood or other abnormal materials. Scraping the 

cut surface of the lung often assists in locating lesions and parasites. 

Figure 10: Pinniped Tracheal Bifurcation IA 

The bronchiolar lymph nodes, 2 to 4 in number, can be found just cranial to the 

heart base adjacent to the aortic arch and pulmonary artery. Incise cross sectionally 

noting any abnormalities. 
HEART: 

The heart of the pinniped is similar to that of a terrestrial mammal, except that 

it is often larger, relative to the animal's size. The pericardium is an opaque sac in 

which the heart rests; this sac is thick walled, and may contain a small amount of 

clear, straw-colored fluid. If the pericardial wall is excessively thickened or there is 

increased fluid accumulation, measure the quantity, assess the quality and 

aseptically take a representative sample and place it in a sterile red top tube. In 

pinnipeds, the myocardium and endocardium (the heart muscle and lining) are 

thicker than in most terrestrial mammals, and are a deep, dark red color. 

Heart dissection in pinnipeds, just as in other mammals, folloys the natural 

physiologic flow of blood, moving from pulmonary vein, to right heart, to lungs, to 

left heart to aorta. To dissect the heart, remove the organ at its atthchment points to 

the pulmonary vein, pulmonary arteries, and aorta. Hold the heaft left (thick) side 

down in the palm of your hand, with the base (the top) of the heark toward the palm 

of you hand and the apex (the pointed end) toward your fingertips. Cut into the 

pulmonary vein and enter the right atrium. Look at the first of fobr heart valves, 

the tricuspid valve, which separates the right atrium from the right ventricle; the 

valve should be opaque, strong, but highly flexible, and of equal d$nsity and smooth 

in texture throughout. In this and all successive valves, note any nodules, 

irregularities or thickened areas. 

Cut into the right ventricle; note any lesions or unusual structqres. Cut up 

through the pulmonary artery (a rather flappy blood vessel that ledves the right 

ventricle at its cranial aspect), noting the pulmonic valve. Next oflen the left 

atrium from the incoming pulmonary artery; the left atrial wall should be thicker 

and more muscular than the right side of the heart. Note the mitrhl valve between 

the left atrium and left ventricle. Enter the left ventricle, which is the thickestwalled of all the chambers of the heart; note any lesions or unusual structures. 

Examine the walls between the right and left atria and right and left ventricles for 

any signs of non-valvular or non-vascular openings between them (these can be 

congenital or hereditary defects, or fetal structures meant to close a4 birth). The 

aortic valve separates the left ventricle from the aorta. The aorta is the most 

muscular artery in the body. Cut through it longitudinally as it exits the heart and 

examine its intimal surface lining for any lesions or unusual structiures. 

As mentioned above, there are four valves in the heart. These are strong, but 

highly flexible, films of tissue, separating the atria from the ventricles and the heart 

from the lungs and aorta. The blood flows into the heart from the pulmonary vein 

at the right atrium, through the tricuspid valve to the right ventricle, to the lungs 

via the pulmonary artery, from the lungs through the pulmonic valve in the left 

atrium, from the left atrium to the left ventricle via the mitral valve and from the 

left ventricle into the aorta through the aortic valve. The thickest muscular tissue 
is in the left ventricle; the thickest-walled artery is the aorta. The most flaccid tissue 

is in the right atrium, and the most flaccid vein the right pulmonary vein. 

THE NERVOUS SYSTEM 

In pinnipeds, the central nervous system (CNS) is composed of the brain and 

spinal cord, both of which lie within bony structures (the skull and spinal 

(vertebral) column). If possible, the brain and spinal cord should be examined on 

every pinniped undergoing necropsy (not those simply exhibiting neurologic signs). 

Before beginning an examination of the CNS, verify any needs for voucher 

specimens. 

The CNS should be removed carefully for closer examinationls. Zt is important 

to realize that the CNS nervous tissue is very fragile, and must be hwdled carefully, 

or it will macerate and disintegrate in your hands. 

I. Cut away and remove the skin, blubber, muscle and connective tissue overlying 

the entire dorsal spine and caudal skull area; 

2. Starting caudally, using a Stryker oscillating saw or cast cutter, lonaitudinally cut 

two lines the entire length of the spinal column into the vertebral bodies 

equidistant (-1 inch to either side) from the animal's dorsal midline. 

3. Lift the tops (dorsal bony covering) off each vertebral body in the spinal column 

exposing the spinal cord beneath. 

4. Move to the base of the skull and again, using a Stryker saw or cast cutter, prepare 

to remove the calvarium (dome-like bony structure that overlies the brain). Use of 

the following three cuts will preserve the skull's bony structure for additional life 

history information. 

Make two symmetrical cuts on each side of the caudal aspect of the skull 

running from each occipital condyle to the foramen magnum. 

Make a third cut perpendicular to the first two cuts transversely aeross the 

frontal bone just posterior to the supraorbital process. 

5. Gently, lift the calvarium off of the brain, gingerly cutting away the brain's 

connective tissue attachments to the inside of the skull with scissors. 

6. Turn the head upside down to allow gravity to assist in gently teasing the brain 

out of the skull. The cranial nerves and connective tissue meninges will have to be 

severed, but then the brain should gently fall onto the table. 
7. Once the brain is out, working from the caudal aspect of the spinal column, 

gently tease the spinal cord out of the vertebral column. You will need to cut 

peripheral nerves and connective tissue to do this, but once you have, you should 

have the entire CNS on the neaopsy table in front of you. 

For ease of viewing you may wish to remove the head prior to brain removal. 

The spinal cord will have to be severed in this process so the entire CNS will not be 

removed intact. Nevertheless to disarticulate the head from the body, at the level 

just caudal to the occipital protuberances make a deep transverse cut down to the 

atlanto-occipital joint (between the skull and the first cervical verteibra), twist and 

disarticulate the skull from the spine. 
3 

----------------------------------------------------- -----------------------------------------------------= 

<EY: 

oc = occipital condyle 

ob = occipital bone 

op = occipital protuberance 

fb = frontal bone 

sc = sagittal crest 

pb = parietal bone 

sw = su~raorbital wrocess 
The cerebrum is the massive, most cranial portion of the brain. The cerebellum 

is caudal to the cerebrum and sits dorsal to the brainstem. The brainstem is at the 

most caudal aspect of the brain, where the majority of the cranial nerves originate, 

and tapers as it moves caudally toward the spinal cord at the first cervical vertebra. 

Examine each structure for symmetry, consistency in color, texture and density, and 

note any bruising, discoloration, constricted or swollen areas, masses, exudates or 

any other abnormalities. Pay particular attention to the pituitary gland found 

protruding from the ventral midline of the brain at the most cranial aspect of the 

brainstem [Figure 121. 

Make cross sectional slices through the brain and spinal cord in a "bread loafing" 

pattern. 

Peripheral nervous system tissues are not normally examined during necropsy. 

If there is a necessity to do so, peripheral nerves are most often located coursing 

alongside major veins and arteries. 
Major Anatomical Divisions of the Brain ..................................................... -----------------------------------------------------= 

KEY: 

c = cerebrum 

cbl = cerebellum 

bs = brainstem 

pg = pituitary gland - 
COLLECTION, PREPARATION AND PRESERVATION 

OF TISSUES AND SPECIMENS 

HISTOPATHOLOGY 

ADDITIONAL EQUIPMENT RECOMMENDED: 

Sharp knife or scalpel blade 

Dissecting tray or cutting board 

Screw top plastic specimen containers containing 10% formalin 

COLLECTION AND PREPARATION OF TISSUES: 

Tissues for histopathology should be obtained with as sharp an instrument as 

possible, preferably a necropsy knife or scalpel blade. In all cases, avoid blunt or 

crushing trauma to the tissues being collected. In areas of observable lesions, be 

certain to take normal as well as abnormal tissue for histopathologic analysis. 

The optimal size tissue sample to yield the best fixation is one agproximately 1 

cm3, although samples as large as 2 cm2 by 0.5 cm are acceptable if adequate fixative 

volume is provided. The container of choice for histopathological specimens is 

plastic, but glass will suffice; the container must have a water-tight svew-top 

closure. Label the specimen jar with an indelible pen, or label the adtual specimen 

directly by inserting aTyvek tag, labeled in pencil, directly into the formalin with the 

specimen. 

If a fetus, with its attendant placental material, is present, take, as aseptically as 

possible, portions of both the fetus and the placenta. Divide these into two sets of 

tissue samples; refrigerate one set, and place the other in 10% neutra/ buffered 

formalin. Immediately contact the laboratory. Maternal blood andbor urine (10 ml 

of each, if available), and fetal fluids, including stomach contents (whatever is 

present), should also be collected and refrigerated. 

PRESERVATION OF TISSUES: 

Standard preservation of tissues and specimens for histopatholoqy is done by 

placing 1 cm3 sections in a container of 10% neutral buffered formalkn (See 

Appendix A). For adequate fixation, the ratio (by volume) of formdlin in the 

container to tissues placed in it must be at least 10:1, i.e. 10 parts formalin to 1 part 

tissue. Follow this ratio religiously, because without adequate fixatiob (caused by 

too large a tissue size or too small a fixative volume) valuable histopathologic 

information will be lost. 
CHECKLIST OF HISTOPATHOLOGIC SPECIMENS: 

This form must accompany any tissues sent for histopathology. 

Specimen Checklist for Histopatholo~y 

Id No. Time/ Date -------- Path Tech 

Blubber Thickness Other Comments: 

Stomach 

- Duodenum 

-Jejunum 

- Ileum - Colon 

- Mesenteric L.N. - Pancreas 

- Spleen 

- Liver 

weight = g 

- Gall Bladder 

- Adrenal, left 

weight = g 

- Adrenal, right weight = g 

- Kidney, left 

weight = g 

- Kidney, right weight = g 

- Urinary Bladder 

- Ovary, right 

weight = g 

- Ovary, left 

weight = g 

- Testis, right weight = g 

- Testis, left 

weight = g 

- Uterus - Cervix 

Vagina 

Prostate 

Penis 

- Diaphragm - Rib 

- Thyroid 

weight = g 

- Parathyroid, left 

weight = g 

- Parathyroid, rt 

weight = g 

- Esophagus - Mediastinal L.N. 

-Thymus 

- Brown Fat 

- Trachea - Bronchus 

- Lung, left 

- Lung, right - Heart 

weight = g 

- Pulmonary Vein - Right Atrium - Tricuspid Valve - Right Ventricle - Pulmonic Valve - Pulmon. Artery - Left Atrium - Mitral Valve - Left Ventricle 

- Papillary Muscle - Aortic Valve - Aorta - Brain 

weight = g 

- Pituitary 

weight = g 

- Spinal Cord 

- Bone Marrow - Eye, left - Eye, right L.N., Periph$ral 

specify___, - L.N., Other 

specify __, - Mammary Gland - Skeletal Mustle - Skin 

- Tongue 

- Tonsils 

Add'l Specimens 
ADDITIONAL EQUIPMENT RECOMMENDED: 

Large and small fecal collection wands 

Glass microscope slides and coverslips 

Containers with 10% formalin 

Forceps 

If the parasite must be sent to a parasitologist for identification-- 

Commercial fecal floatation set-up 

Physiological saline 

Petri dishes 

Warm 70% ethanol 

Phenol 

Absolute alcohol 

AFA 

Solution of 2.5% potassium dichromate 

COLLECTION, PREPARATION AND PRESERVATION OF TISSUES: 

The study of parasites, even though many may be observed while examining a 

pinniped, is still a very specialized field. For routine fecal floatation on-site, collect 

4-5 grams of feces from the terminal colon. If any delay in examinatjon or 

preparation is anticipated, the sample should be placed under refrigeration (4" C). 

Except for hookworm eaG which deteriorate rapidly, most parasite eggs can remain 

identifiable for days if the fecal sample is kept refrigeratedlg. In the case of coccidial 

or~anisms, place 1 part feces in 5 parts of potassium chromate in a jag and package 

for shipping. 

To prepare a blood specimen for parasitological examination, collect I ml of 

blood. Place one drop on a clean, glass microscope slide. Place the regt in a red top 

tube and add 9 ml of 10% neutral buffered formalin. All parasites, whether from 

feces or blood, if fixed in-house, should be mounted on microscope slides using 

piccolyte sealant under the coverslip as the mounting media. Label all specimens. 

It is not unusual for the layperson, volunteer, technician or veterinarian to be 

unable to identify the genus and/or species of a parasite being collected. If a fecal 

sample is to be sent for parasitological examination, collect 5-10 grams of feces and 

place in a container with 10% neutral buffered formalin at a ratio of 31 

forma1in:feces. 
For preparation of an individual parasite specimen, if the parasite is particularly 

dirty, first gently wash it with a syringe-full of physiological saline. Then, collect the 

parasite and the tissue to which it may be attached, and place it in a jrar containing a 

9:l solution of 10% formalin:tissue, applying a tightly fitting lid. 

For clearing nematodes, briefly place the parasite in a petri dish containing 

warm 70% ethanol, until the specimen relaxes and stretches out straight. Next flood 

with a 4:l mixture of pheno1:absolute alcohol, and leave the petri di$h uncovered in 

a dust-free area until all the liquid has evaporated. Store the specimen in 70% ethyl 

alcohol. 

For clearing cestodes or trematodes, place the individual parasite (if it can be 

isolated) in a petri dish of physiologic saline in the refrigerator (4" C) for 2 hours to 

relax the specimen and allow it to release any eggs if present. Once t&e parasite is 

relaxed, take a teasing needle and gently straighten out the parasite. Preserve it and 

its eggs, if present, by slowly flooding the petri dish with a hot (42-52@ C) solution of 

AFA (see Appendix A); allow it to sit for a minimum of 1-2 hours (overnight is 

optimal). Store the specimen in 70% ethyl alcohol. 

Label the specimen, including the host animal's (pinniped's) species, age and sex. 

Note where the parasite was located, and any gross pathological changes associated 

with it, and include this information with the specimen. 

ADDITIONAL EQUIPMENT RECOMMENDED: 

Sterile surgical gloves in appropriate sizes 

Alcohol lamp or Bunsen burner and matches/lighter 

New, sterile #15 and #10 scalpel blades with handle 

Sterile swabs or culturettesf 

Sterile red top tubes 

Refrigeration 

COLLECTION AND PREPARATION OF TISSUES: 

If bacterial disease is suspected, because motile intestinal bacteria can seed to 

many organs within 5 to 10 minutes after death, it is best to minimize 

contamination of the sample by collecting bacterial samples as soon as the carcass's 

abdomen or thorax is openeds. 
It is essential that material to be cultured for bacteria be collected in a sterile 

manner. First, check to make certain there are no flammable chemicals in your 

work area. Then, light the alcohol/Bunsen burner. Heat your #15 (sterile, sharptipped) scalpel blade in the flame for approximately 15 seconds. Immediately bake a 

stab incision into the organ you wish to culture. The hot blade will singe the tissue, 

killing any organisms present on the tissue's surface. Taking precautions not to 

contaminate (with hands, coughing, non-sterile instruments) the area of interest, 

carefully insert a sterile swab or culturette into the stab incision. Gently roll the 

swab or culturette around inside the incision. Carefully remove the swab and 

immediately place it (without touching anything, including the outside of your 

transport container) in a sterile container. The sterile container may be one 

provided by your laboratory, one supplied with the culturette, or a sterile red top 

tube ordinarily used for blood collection. 

If pus (purulent liquid material) is present, using a sterile syringe, collect flhe 

material leaving no air bubbles in the syringe. This will allow for culture of 

potential anaerobic (non-oxygen needing) bacteria at the laboratory. 

If a bacterial infection is suspected in a solid organ, such as the liver, kidney or 

lung, sterilize your scalpel blade in the alcohol/Bunsen burner flame, and cut a 

wedge of tissue, approximately 2 to 4 cm square, which includes the suspect ariea, 

and place the sample in a Whirl-Pak bag or a red top tube. If an intestinal bacterial 

infection is suspected, use string to tie off both ends of a small section (about 2 cm) of 

intestine, cut each side of the section beyond the tie-off pointed and place the tubelike specimen in a Whirl-Pak bag. If a brain infection is suspected, as aseptically as 

possible, remove the skull with a Stryker saw or rongeurs, and aseptically take a 3 

cm2 piece, placing it directly in a Whirl-Pak bag or sterile red-topped tube. Label any 

specimen collected. 

PRESERVATION: 

Any bacteriological sample must be stored under refrigeration (DO NOT Rreeze). 

Immediately request a laboratory pick-up or personally before the end of the day, 

take the material td the laboratory yourself. Never allow a bacteriological sar$ple to 

sit overnight unattended! It is best to allow the laboratory the opportunity to plate 

the bacterial sample within 4-6 hours, since some bacteria will die if left overnight at 

4" C. 
In general, the size, location and preparation of specimens to be submitted24 for 

biotoxicologic examination are as follows: 

Stomach contents: -1 /2 frozen (-20" C) 

-1 /2 refrigerated (4" C) 

-10 grams in equal volume of 10% 

buffered formalin (for algal identification)25 

Liver: 100 grams Left caudal lobe Frozen 

Kidney: 100 grams Left caudal pole Frozen 

Blubber: 100 grams Ventral Midline Frozen 

(just cranial to the xiphoid proces$) 

Place each of these specimens in a clean, plastic container, with a tight fittjng top. If 

the animal's size or carcass condition precludes specimens of the recommended size 

above, take whatever is available. 

Toxic algal bloom material is most often found in stomach contents, whether 

free or incorporated into food materials, such as fish. So stomach contents are the 

most valuable material for diagnosis of biointoxication; accompanying swples of 

liver, kidney and blubber may be helpful to the toxicologist in correlating data. Even 

if biotoxins are strong suspects, always send in tissues for histopathology as well, and 

be certain to inform the toxicologist that histopathology is pending. 

Liver, kidney and blubber once collected, should be frozen at 

-20' C, in a standard walk-in or chest freezer. The freezer should not be "frost-free", 

as is found in refrigerator-freezer combinations and some chest freezers, because 

some toxins are sensitive to degradation by the constant freeze-thaw cycleb that 

materials in a frost-free freezer undergo. 

HEAVY METALS AND ANTHROPOGENIC (man-made) ORGANICS: 

ADDITIONAL EQUIPMENT RECOMMENDED 

Teflon sheeting (for clean working surface) 

Teflon jars with'lid labels (if unavailable, acid rinsed glass jars covered with 

plastic film prior to replacing lid for heavy metals and acetone rinsed jars 

covered with acetone rinsed aluminum foil prior to replacing the lid for 

organics) 

Stainless steel knives 

Titanium knife with Teflon handle 

High grade ethanol and distilled water for rinsing knives 

Whirl-pak or zip-lok plastic bags (for heavy metal specimens) 

Heavy duty aluminum foil (for organic specimens) 
COLLECTION AND PREPARATION OF TISSUES 

Because of the possibility that presence of contaminants may lead to litigation 

and because analyses may involve detecting extremely small amounts of 

contaminants, utmost care must be taken in the collection and proce~sing of tissues. 

The chain-of-custody procedures detailed under Shipping of Tissues and Specimens 

should be followed. Ideally, specimens should be collected under canditions that 

avoid contamination. Avoid salt water, tobacco smoke, vehicle exhausts, bug sprays 

and other aerosolized foreign materials during collection. Recognizing that a 

laboratory and special materials such as Teflon containers and titanium knives are 

not always available, if care is taken, specimens can be collected with ethanol-rinsed 

stainless steel instruments and subsampled by the laboratory performing the 

analyses. 

The tissues and other specimens routinely collected from pinnip~eds for toxicant 

analyses are blubber, liver, kidney, brain; and for special considerations, urine, 

stomach contents, blood serum, whole blood, reproductive organs, milk, and feces. 

The size of specimens to be collected, if available, is: 

Blubber 

Liver 

Kidney 

Brain 

Urine 

Stomach contents 

Seru,m 

Whole blood 

Milk 

300 grams 

300 grams 

300 grams 

300 grams 

500 ml. 

500 ml. 

50 ml. (in red top tube) 

50 ml. (in purple top tube) 

50 ml. (in red top tube) 

It is important that toxicological findings are correlated with geaeral signs and 

lesions. Therefore, gross necropsy reports should accompany any specimens, and 

histopathological results, if any, should be received as soon as they $re completed. 

General: 

Tissue samples are to .be collected as rapidly as possible after opehing body cavity 

in order to prevent contamination add deterioration. 

Stainless steel knives should be used to make initial opening of the body cavity. 

(Do not use titanium knives to cut through skin since this dulls the knife. Only 

use them for collecting the actual specimen material). 
Blades should be rinsed with ethanol before a tissue sample is taken. Because 

ethanol will fix blood on a metal surface, blades should be rinsed with distilled 

water first and then ethanol after each sample is taken and before a new sample 

is collected. 

Except where indicated above, specimens should be placed in Teflon bags or 

containers. If such containers are not available, the specimen shlould be divided 

and placed into separate containers for heavy metals or organics (See Additional 

Equipment Recommended). If the appropriate containers are nQt available, the 

specimens should be divided and wrapped in plastic wrap (for heavy metals) or 

heavy duty aluminum foil (for organics). 

All specimens (except whole blood) should be frozen immediataly after 

collection. Ideally, each sample should be frozen by immersion in liquid 

nitrogen for 10 minutes or frozen at -70°C or lower. If such super-cold 

equipment is not available, samples should be frozen at -20°C. At a minimum, 

refrigeration is required until a dry ice source or freezer (home chest freezer or 

walk-in freezer only) can be located. A+refrigerator's freezer is not adequate for 

long-term storage. Freezing to very low temperatures is not necessary for tissues 

awaiting heavy metal analysis. 

Individual Tissues--Instructions 

Blubber 

Collect on ventral midline just cranial to the 

xiphoid process. 

Separate blubber from underlying muscle layer and 

skin. 

Wash tissue in high grade distilled distilled water to 

remove blood and other fluid before freezing. 

Liver 

Collect 150-200 gram section from each side of the liver at 

the distal lobes on their medial aspect. Wash tissue in high grade 

distilled water to remove blood and other fluid before freezing. 

Kidney 

Collect from left caudal pole. 

Wash tissue in high grade distilled water to remove blood and other 

fluid before freezing. 
Brain 

Halve longitudinally. 

Whole Blood 

Should not be frozen. Refrigerate for immediate analysis. 

SHIPPING AND LABORATORY ANALYTICAL TECI-INIQUES 

If a liquid nitrogen shipper is available, it should be used. The liquid nitrogen 

shipper should be filled with liquid nitrogen for at least six hours to fully prepare it 

for shipping. Pour off excess liquid nitrogen and place the frozen samples in the 

shipper. 

If a liquid nitrogen shipper is not available, specimens should be shipped under 

conditions to avoid thawing. Specimens should be placed in an insulated container 

with an adequate amount of dry ice to prevent thawing. 

Whenever a shipment is made, the receiving laboratory should be notified in 

advance of shipment. Unless special arrangements are made, specimens should not 

be shipped over a weekend. Documentation describing the animal, gross necropsy 

results, and other information should be included. 

Under certain circumstances, e.g., during an unusual mortality event, 

individuals may be given specific instructions as to which laboratories should 

perform analyses and what analyses should be performed. If analysds are being 

conducted for an independent investigator, the laboratory must be given specific 

instructions on the type of analyses to be performed. A statement such as "analyze 

for toxic substances" is entirely inadequate. 

It is essential to check with the toxicology laboratory to be certain that it is 

capable of conducting the test(s) desired. Make certain that the laboratory chosen is 

staffed by professional analytical chemists, pathologists, and techni~i~ans, regularly 

trained and up-to-date in the use of modern instrumentation and the sophisticated 

techniques toxicological analyses required. 

Laboratory analytical methods are continually being changed and improved. 

Laboratories should have documentation of proper validation, inclyding standards 

curves for standard reference materials (SRMs), revalidation of analytical methods 

(standard operating procedures), documentation of method accuracy and precision, 

and a quality assurance program. 
Such validation allows a greater degree of standardization and interlaboratory 

reliability. In addition, it is important to realize that every chemical analysis 

performed in a laboratory has both a limit of detection and a (higher) limit of 

reliable quantification associated with it. The researcher or cliniciao should know 

what these limits are, not only for the compound itself, but also for the matrix 

within which the compound exists, e-g., liver, blubber, etc. A toxicdlogy report from 

a laboratory of "not present in detectable amounts" is generally appropriate, whereas 

a report of "not present" is generally inappropriate. Levels of detection must be 

given for the compound. 

The laboratory must have a written protocol in place for each as$ay, as well as 

documented quality control procedures, not only of analytical tech+ques and 

methods, but also for personnel performing analyses. If there is any suspicion that 

the data from materials you are sending to a laboratory for analysis iwill be needed in 

a court of law, be certain the laboratory's personnel are trained and regularly 

updated regarding instrumentation and procedures for which they have 

responsibility. Make certain that the laboratory records processing errors, equipment 

failures, or quality control standards outside the norm. These are the most reliable 

ways to prove that a laboratory is scientifically credible. 

In cooperation with the National Marine Fisheries Service, the National 

Institute of Standards and Technology has a marine mammal quality assurance 

program in place that allows for interlaboratory comparative exerci$es, standardized 

calibrations, and the provision of SRMs. The American Associatioe of Veterinary 

Laboratory Diagnosticians (AAVLD) and the College of American Pathologists (CAP) 

also accredit many private and public laboratories in the United Staltes. 

It is also helpful to choose a laboratory which has rapid reporting, by telephone 

or FAX, followed by transmission of original documentation by mdl. Data received 

from a laboratory regarding any test performed on tissues from an animal must be 

kept in that animal's permanent case record folder. If it is anticipated that litigation 

may be involved, the case folders and all specimens from the case must be kept 

under lock and key, and a strict chain of custody kept on the recordt;. 

MISCELLANEOUS DIAGNOSTIC SPECIMENS 

Always be certain prior to proceeding that you contact the diagnostic or research 

laboratory of choice to insure that appropriate personnel, equipment and supplies 

are on hand for performing the special test analysis you are requesting. 
Isozyme studies require the collection of blood or tissue. Collect whole blood in 

sodium citrate (blue top tube), centrifuge for 10 minutes at 10,000 rptns, draw the 

plasma off the top of the tube and send on ice immediately or freeze until sent to the 

laboratory. A minimum of 10 to 100 grams of muscle tissue can also be aseptically 

collected and refrigerated or placed on ice to await immediate shipment to the 

laboratory. 

CYTOCHROME P-450:27 

Tissues should be as fresh as possible, preferably collected withi0 30 minutes 

following death, since they will be used for analyses of halogenated and polynuclear 

aromatic hydrocarbons. Liver, lung, heart, kidney, small intestine, and skin samples 

of 100 grams each may be collected. Tissues should be divided into $0 gram 

allotments, each folded into a piece of aluminum foiled and labeled inside and out. 

These samples are then frozen, preferably in liquid nitrogen, or placed on dry ice for 

immediate transport to the laboratory of choice. Be certain to call the laboratory 

ahead of time to inform them of the pending shipment. 

DNA STUDIES:28 

Blood in the live pinniped, or heart muscle, skin, sperm or skelletal muscle from 

the freshly dead pinniped, are adequate to perform DNA isolation. In the live 

animal, collect a minimum of 10 m of blood in an EDTA (purple top) or sodium 

citrate (blue top) tube. Once collected, place in a refrigerator (at 4O C) or on ice for 

pick up by the laboratory. If the blood must wait for pick up, it is best to centrifuge it 

for 5 minutes, then perform two steps: (1) decant (pour or draw off) the straw 

colored plasma, label it and freeze it, and (2) pipette off the buffy codt (I to 3 mm 

whitish layer between the red blood cells and plasma), place it in a deparate vial, and 

label and freeze it. Refrigerate (DO NOT FREEZE) the red blood cells. Label the 

specimen and contact the laboratory. 

In the freshly dead pinniped, collect heart or skeletal muscle, skin, gonadal 

tissue, or sperm. The tissue sample size to take is approximately 2 to 10 grams; this 

material should be sliced with a sharp, clean knife or scalpel blade into 1 cm thick 

sections prior to fixing. To fix, either freeze at -20 to -70° C, or preserve in 70% 

ethanol or isopropyl alcohol. 
TOOTH/ BONE /SKELETAL PREPARATION 

ADDITIONAL EQUIPMENT RECOMMENDED: 

Nylon mesh bag 

Large barrel 

or 

3 foot deep hole in the ground 

or 

flesh-eating beetles 

Trisodium phosphate 

Large Pot 

Stove 

COLLECTION AND PREPARATION OF SKELETAL REMAINS: 

Once necropsy examination is complete, if the skeleton is needed for natural 

history studies, remove as much skeletal muscle as is possible from the skeleton and 

place it in a mesh bag. Either bury the bag at least 3 feet under the soil surface for 4 

weeks, or place in shallow ocean (attached to a point on shore) for 2 to 5 days. 

Other methods involve placing the skeleton in a specially prepareH 50 gallon 

drum containing flesh-cleaning beetles for up to 4 weeks. Alternatively, the bone, 

may be placed in a detergent solution for approximately two weeks to de-grease it, 

followed by a weak ammonium hydroxide solution for 1-2 days maxihnum (to 

bleach the bones)29. Once cleaned, bony materials can be frozen, placed in formalin, 

or air-dried, depending on their intended use. 

TOOTH PREPARATION FOR AGING PINNIPEDS: 

In order to age a pinniped, the best method is to extract a canine tooth from the 

upper jaw. In the field, the tooth, once collected, can biz refrigerated, frrozen or 

packed in salt30. At an institution, boil the tooth in a solution of trisodiurn 

phosphate (diluted at 1 Tablespoon/quart of water) for approximately 15 minutes to 

remove the pulp. Then air dry the tooth. Once dry, using a high intensity light and 

hand lens (or a dissecting microscope), the horizontal growth rings of the tooth can 

be counted. A pair of rings (one "light" and one "dark") equals 1 year of age. Some 

institutions cut the tooth longitudinally for more ease in counting rings. 
PACKING MATERIALS: 

Each specimen's container should be clearly labeled with indelible ink, including 

date collected, species, sex, identification number, source of tissue/body fluid, and 

the collector's name or institution. If being shipped for analysis, the tibsues should 

be accompanied by a copy of the gross necropsy report. If a number of specimens are 

to be shipped in the same package, label the package inside and out with your name 

and address (the sender) and the name and address (the receiver) to which the 

package is being shipped. 

Prepare a form in quadruplicate [See Appendix C] listing each specimen, the date 

collected, species, sex, identification number, source of tissue/body fluid and to 

whom the specimen is being shipped. Enclose one copy inside the package, place 

one in an envelope addressed specifically to the laboratory of choice add attached 

firmly to the outside of the shipping container, send one form with yOur monthly 

report to your Regional National Marine Fisheries Service Office, and keep one on 

file, attached to the necropsy report at your facility. Use smear-proof ihk to insure 

legibility. 

Each time a tissue sample or specimen leaves your facility, it must be 

accompanied by its own signed Chain of Custody Form [See Appendix Dl. Sign next 

to the same printed name and include the Form with the specimen. Each person 

who handles the sample for whatever reason must sign and date the dorm as noted 

above. 

Samples should be tightly packed to avoid jostling, breakage, or thpwing during 

shipment; newspaper or butcher paper assists in insulation during shi ping. 

Shipping of medical diagnostic specimens by air or mail is subject to F 1 deral 

regulations. Rules require proper packaging, including securely sealin the package, 

adding sufficient absorbent materials to take up the contents in case of C leakage, and 

including shock resistant material to withstand ordinary handling during transitsl. 

The outside of the package should be labeled with "DIAGNOSTIC SPHCIMEN" 

written in large identifiable letters. 

The importance of communication between the sender and receiver cannot be 

overemphasized, particularly if the specimens being shipped are valuable biological 

and/or legal samples. 
SHIPMENT OF CLINICAL PATHOLOGICAL SPECIMENS: 

Shipment of blood and bodily fluids collected from live (or very recently dead) 

pinnipeds, must be shipped to the laboratory as quickly as possible following 

collection. Have a carrier or courier service pick up and deliver the specimens to 

the laboratory or airport; if possible, do this yourself or direct a responsible person to 

do this for you. Emphasize the importance of rapid delivery of samples. For 

transport via air or over long distances, samples should be kept cool, by packaging 

them in a styrofoam container. Serum can be shipped frozen and pracked on dry ice. 

SHIPMENT OF HISTOLOGICAL SPECIMENS: 

Histopathological specimens for shipment should be well-sealea (a piece of 

packing tape around the container's screw top will help prevent leakage). If large 

numbers of tissues must be shipped, tissues in 10:1 formalin:tissue, once entirely 

fixed (3-4 days), may be removed and placed in smaller amounts of 10% buffered 

formalin for shipping purposes. 

SHIPMENT OF MICROBIOLOGICAL SPECIMENS: 

Bacteriological and mycological specimens should be kept chilled during 

transport; ship in styrofoam containers and include at least two or tthree blue-ice or 

similar frozen gel packs. If transportation time is expected to be looger than 24 

hours, the samples must be packed in dry ice. Viral specimens mu$ be frozen; 

frozen specimens should be maintained while being shipped by using an insulated 

container made of styrofoam and packed with dry ice. Make certain the styrofoam 

container is well-sealed with packing tape to keep the cold in during transport. 

SHIPMENT OF PARASITOLOGICAL SPECIMENS: 

It is best to keep parasitological specimens cool during shipment. Double bag, or 

seal screw top containers with packing tape, to prevent leakage, and ship in 

styrofoam containers if possible. 

SHIPMENT OF TOXICOLOGICAL SPECIMENS: 

Toxicological specimens must be shipped frozen. Place them in styrofoam 

containers with either dry ice or multiple frozen gel packs. Make certain the 

styrofoam container is well-sealed with packing tape and/or enclosed in a cardboard 

packing box to keep the cold in. 
CONCLUSIONS 

The importance of history, findings, and lesions must not be underestimated 

in shipping specimens to a diagnostic laboratory. Such information (such as a 

complete history, a good description of signs, and an accurate descripltion of gross 

lesions) is invaluable, not only to the clinician or diagnostician, who must 

intelligently choose appropriate tests to be run by the laboratory, but also to the 

laboratory itself, which must determine the diagnostic methodology to be used and 

the significance of pertinent results. Each of these steps is a piece of hhe puzzle, a, 

final diagnosis [Figure 131. 

The quality of tissues and other specimens collected either at ar prior to death 

has more to do with an accurate diagnosis than history or clinical finldings. Since in 

pinnipeds, history and clinical findings are often missing, tissue and specimen 

collection becomes even more crucial. Without proper handling, cdllection, 

preparation, preservation and shipping, valuable data, and scientific evidence 

admissible in a court of law, may be lost forever, and a legally defensible diagnosis 

never defined. 

In addition, tissues shipped to accredited laboratories that follcpw quality 

assurance and quality control procedures and accomplish this through good 

laboratory practices, help insure that the diagnostic laboratory data generated will be 

of greatest use. 
The main purposes in producing this Pinniped Forensic Guide are: 

to encourage standard operating procedures for field and necropsy room 

investigations of pinniped strandings and unusual mortality events; 

to assist in producing standard reference materials for comparative studies; 

to recommend analytical protocols that yield comparable diagnosec results; and, 

to improve the overall quality of tissue and specimen collection during pinniped 

beachings, strandings and unusual mortality events, so that a clinidal diapnosis of 

the cause of an event mav hold up in a court of law should such didgnosis be 

challenped. 

Every specimen in every container in every styrofoam cooler in every 

shipment, must be considered as the most important specimen, and1 treated as if it 

alone could be the specimen that proves or disproves a case. 

Anyone and everyone can perform the procedures in each Sction of this 

Guide. The overall goal of this guide is to provide consistency in collection of 

information for use in establishing a disease or mortality event dia nosis. 

Information obtained from proper and dependable procedures yiel d s the most 

effective clinical and laboratory diagnostic data, which in turn cont@ibutes to 

tentative and final diagnoses. The information obtained also furthers the known 

body of science on the particular species you are investigating, and provides 

reference materials for use in future studies and/or mortality events. Skillful 

techniques, with proper specimen and tissue collection and preservation, yield 

maximum reliability and credibility of scientific information. 

Even if you have little or no experience with marine mammhls, you can still 

offer significant value to the collection of scientific data, because th~ key to credible 

information is describing what you are seeing rather than what you think you 

should be seeing. Accurate observations and simple descriptive tetms are the 

foundations upon which the veterinarians and researchers who are required to 

provide a final diagnosis as to cause of death base their decisions. 

The more geographically isolated or remote you are from reqdily accessible 

clinical laboratory facilities, the more carefully you will need to obsprve and collect 

data to yield the most accurate and usable of diagnostic tissue/specimen samples and 

assertions. Never omit the simple diagnostic procedures and move directly to the 

more complex procedures, because this is ignoring the big picture, where subtle 

hints and changes may yield the most surprising details. 
Always remain aware that the information you are collecting and recording is 

useful in multiple and meaningful ways. Not only can such data sefie as an early 

warning system, cuing researchers and the public to potentially larger problems 

hidden in the background, but also as materials and data able to be fully admissible @ in a court of law to prove beyond a reasonable doubt that what the diagnostician 

interprets as the cause of an event can be argued before and interpreted by a judge or 

jury. 
APPENDIX A 

Formulations for Commonlv Used Fixatives 

10% Neutral Buffered Formalin: 

Formaldehyde solution (37-40%) 100 ml 

Distilled water 900 ml 

Sodium acid phosphate, monohydrate 4.0 grams 

Disodium phosphate, anhydrous, pH=7.3 6.5 grams 

AFA (Alcohol-Formalin-Acetic Mixture): 

Formalin (as above) 4 ml 

Ethanol (95%) 20 ml 

Glacial acetic acid 0.5 ml 

Distilled water 16 ml 
APPENDIX B 

Laboratories Accredited bv the 

American Association of Veterinary Laboratory Dia~nosticians -, 

July, 199232 

1. County of Los Angeles, Department of Health Services 

Division. Comwrative Medical and Veterinarv Public 

Health Skrvice; Downey, CA , 

2. Colorado Veterinary Diagnostic Laboratory, College of 

Veterinary Medicine, Colorado State University, 

Fort Colfins, CO 

3. Department of Pathobiology, University of Connecticut, 

Storrs, CT 

4. Florida Veterinary Medicine Diagnostic Laboratory 

System, Florida Department of Agriculture, Kissimmee, FL 

5. The Division of Comparative Pathology, University of 

Miami, Miami, FL 

6. Diagnostic Assistance Laborato College of Veterinary 

Medicine, University of Georgia, ~zens, GA 

7. Veterinarv 1)iaenostic and Investieational Laboratom. , . College of ~eterin'a'ry Medicine, uni<k'rsity of Georgia, 

Tifton, GA 

8. Laboratories of Veterinary Diagnostic Medicine, College 

of Veterinary Medicine, University of Illinois, Urbana, IL 

9. Illinois Department of Agriculture, Animal Disease 

Laboratory, Galesburg, IL 

10. Illinois Department of Agriculture, Animal Disease 

Laboratoryr, Centralia, IL 

11. Indiana Animal Disease Diagnostic Laboratory and 

Southern Indiana Purdue Agricultural Center, Purdue University, West Lafayette, IN 

12. Iowa Veterinary Diagnostic Laboratory, College of 

Veterinary Medicine, Iowa State University, Ames, IA 

13. Kansas Veterinary Diagnostic Laboratory, Department 

of Veterinary Diagnostics, College of Veterinary Medicine, 

Kansas State University, Manhattan, KS 

14. Murray State University Veterinary Diagnostics and Research Center, Hopkinsville, KY 

15. Louisiana Veterinary Medical Diagnostic Laboratory, School of Veterinary Medicine, LSU, Baton Rouge, LA 

16. The Division of Diagnostic Laboratories, School 

Veterinary Medicine, Tufts Univetsity, North Grafton, MA 

17. Animal Health Diagnostic Labbratory, College of Veterinary Medicine, Michigan State University, East Lansing, MI 

18. Minnesota Veterinary Diagnostic Laboratory, College of 

Veterinary Medicine, University df Minnesota, St. Paul, MN 

19. Mississi pi Veterinary Diagnostic Laboratory, 

~ississippi bepartment of Agricdturq Jackson, MS 

20. Veterinary Medical Diagnostiq Laboratory, College of 

Veterinary Medicine, University of Missouri, Columbia, MO 

21. State of Montana Animal Health Division, Diagnostic 

Laboratory, Bozeman, MT 

22. Nebraska Veterinary Diagnostic Laboratory System, 

Department of Veterinary Science, University of Nebraska, 

Lincoln, NE 

23. Veterinary Diagnostic Laboratory, New York State 

College of Veterinary Medicine, Cornell Univ, Ithaca, NY 

24. North Carolina Veterinary Medical Diagnostic Laborato System, North Carolina Department Agriculture, 

Raleigh, % 

2.5. North Dakota State Veterinary Diagnostic Laboratory, 

Department of Veterinary Scienceu North Dakota State Univ, 

Fargo, ND 

26. Oklahoma Animal Disease Dl gnostic Laboratory, College of Veterinary Medicine, klahoma State University, 

Stillwater, OK 

8 

27. Animal Disease Research and Diagnostic Laboratory, South Dakota State University, Btookings, SD 

28. & 29. Texas Veterinary Mediqal Diagnostic Laboratory, 

Texas A&M Univ System, Amarillo, TX & College Station, TX 

30. Washin on Animal Disease Diagnostic Laboratory, 

College of P eterinary Medicine, Washington State 

University, Pullman, WA 

31. Central Animal Health Laboratory, Wisconsin 

Department of Agriculture, Mineral Point Road, Madison, WI 

1 32. Wyoming State Veterinary Laboratory, Department of 
APPENDIX C 

Sample Transfer Record 

Project Name Project No. ---------------- 

Date - - - Time (24 hour clock) 

day mo Yr 

Released By: ................................. ID No --d------------ 

name 

location 

Date - - - Time (24 hour clock) 

day yr 

Received By: ................................. ID No. -------------- 

name 

location 

Comments: 
Marine Forensics 

& Protected Resources 

Program -- 

Chain of Custody Form 

Sample descript 

Every penon subsequently In possession of the sample must complete a rece~pt/release box on the back of this 

form. (Each person must sign and date the form twrce, once for receipt of the sample, and once for release). 
- 

Rece~pt scgnature Rece~pt date 

Pr~nt narnelagency 

Release signature. Release date: 

Method of transfer to next person' 

J 

Receipt signature. Receipt date. 

Print narnelagency: 

Release signature: Release date: 

Method of transfer to next person: 

Receipt signature: Receipt date: 

Print namelagency: 

Release signature: Release date; 

Method of transfer to next person: 

Receipt signature: Receipt date: 

Print namelagency: 

Release signature: Release date: 

Method of transfer to next person: 

1 

Receipt signature: Receipt date: 

Print namelagency: 

Release signature: Release date: 

Method of transfer to next person: 

Receipt signature: Receipt date: 

Print namelagency: 

Release signature: Release date: 

Method of transfer to next person: 

Receipt signature: Receipt date: 

Print narnelagency: 

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REFERENCES 

I. Roletto, J. and Mazzeo, J., 1990. Identification of North American Marine 

Mammals, in Handbook of Marine Mammal Medicine: Health, Disease and 

Rehabilitation, L.A. Dierauf (ed.), CRC Press, Boca Raton, FL, pp. 399-468. 

2. Geraci, J.R. and Lounsbury, V.J.,1993. Pinnipeds, in Marine Mammals Ashore: 

A Field Guide for Strandings, A Texas A & M Sea Grant Publication, Galveston, 

TX, pp. 58-69. 

3. Wilkinson, D. M., 1991. Report To: Assistant Administrator for Fisheries, 

Program Review of the Marine Mammal Stranding Networks, National Oceanic 

and Atmospheric Administration, Appendix B. 

4. Geraci, J.R. and Lounsbury, V.J., 1993. Specimen and Data Collection, in Marine 

Mammals Ashore: A Field Guide for Strandings, A Texas A & M Sea Grant 

Publication, Galveston, TX, p. 179. 

5. Dierauf, L.R., 1990. Pinniped Blood Collection, in Handbook of Marine Mammal 

Medicine: Health, Disease and Rehabilitation, L.A. Dierauf (ed.), CRC Press, Boca 

Raton, FL, pp. 582-584. 

6. Geraci, J.R. and Lounsbury, V.J., 1993. Specimen and Data Collection, in Marine 

Mammals Ashore: A Field Guide for Strandings, A Texas A & M Sea Grant 

Publication, Galveston, TX, p. 179. 

7. Williams, T.D., Williams, A.L. and Stoskopf, M., 1990. Marine Mammal 

Anesthesia (Pinnipeds), in Handbook of Marine Mammal Medicine: Health, Disease 

and Rehabilitation, L.R. Dierauf (ed.), CRC Press, Boca Raton, FL, pp. 183-188. 

8. Bossart, G.D. and Dierauf, L.R., 1990. Marine Mammal Clinical Lalboratory 

Medicine, in Handbook of Marine Mammal Medicine: Health, Disease and 

Rehabilitation, L.R. Dierauf (ed.), CRC Press, Boca Raton, FL, p. 47. 
9. Spraker, T.R, 1985. Basic Necropsy Procedures, in Clinical Textbook for Veterinary 

Technicians, D.M. McCurnin (ed.), W. B. Saunders, Philadelphia, pp. 483-484. 

10. Sweeney, J. C., 1990. Surgery, in Handbook of Marine Mammal Medicine: Health, 

Disease and Rehabilitation, L.R. Dierauf (ed.), CRC Press, Boca Raton, 

FL, pp. 222-224. 

11. Cowell, R.L. and Tyler, R.D., 1989. Cytology of Cutaneous Lesions, in The 

Veterinary Clinics of North America: Small Practice, vol. 19, no. 4, July, B.W. 

Perry (ed.), W.B. Saunders Co., Philadelphia, PA, pp. 769-775. 

12. National Marine Fisheries Service, Southeast Fisheries Center, Charleston 

Laboratory, 1993. Forensic Manual (for the Analysis of Marine Animal Tissues), 

[Draft], p. 42. 

13. Dierauf, L.R., 1990. Aging the Pinniped via Dentition, in Handbook of Marine 

Mammal Medicine: Health, Disease and Rehabilitation, CRC Press, Boca Raton, FL, 

p. 559. 

14. Buck, C.D. and Schroeder, J.P., 1990. Public Health Significance of Marine 

Mammal Disease, in Handbook of Marine Mammal Medicine: Health, Disease 

and Rehabilitation, CRC Press, Boca Raton, FL, pp. 163-174. 

15. Strafuss, Albert, C., 1988. Descriptive Writing of Pathologic Changes, in Necropsy: 

Procedures and Basic Diagnostic Methods for Practicing Veterinarians+ A. Strafuss 

(ed.), Charles C. Thomas, Publisher, Springfield, IL, pp. 43-49. 

16. Strafuss, Albert, C., 1988. Removal of the Brain and Spinal Cord, in Necropsy: 

Procedures and Basic Diagnostic Methods for Practicing Veterinarian9,A. Strafuss 

(ed.), Charles C. Thomas, Publisher, Springfield, IL, pp. 30-32. 

17. Dierauf, L.R, 1990. Marine Mammal Parasitology, in Handbook of Marine 

Mammal Medicine: Health) Disease and Rehabilitation, CRC Press, Boca Raton, 

FL, pp. 89-90. 
18. Brown, H. W. and Neva, F.A., 1983. Technical Diagnostic Methods, Basic Clinical 

Parasitology, Appleton-Century-Crofts, Norwalk, CT, p. 315. 

19. Dunn, J. L., 1990. Microbial Sampling Techniques, in Handbook of Marine 

Mammal Medicine: Health, Disease and Rehabilitation, L.R. Dierauf (led.), CRC 

Press, Boca Raton, FL, pp. 73-74. 

20. Kennedy-Stoskopf, S., 1990. Immunology of Marine Mammals, in Handbook of 

Marine Mammal Medicine: Health, Disease and Rehabilitation, L.R. Dierauf, (ed.), 

CRC Press, Boca Raton, FL, pp. 115-124. 

21. Kennedy-Stoskopf, S., 1990. Virus Isolation--An Overview, In Handbook of 

Marine Mammal Medicine: Health, Disease and Rehabilitation, L.R. Oierauf (ed.), 

CRC Press, Boca Raton, FL, pp. 97. 

22. National Marine Fisheries Service, Southeast Fisheries Center, Charleston 

Laboratory, 1993. Forensic Manual (for the Analysis of Marine Animal Tissues), 

[Draft], p. 60. 

23. Leland, D. S., 1992. Collection and Transport of Specimens, Laboratory Diagnosis 

of Viral Infections, 2nd Edition, E. H. Lennette (ed.), Marcel Dekker, Ific., New York, 

pp. 3-43. 

24. Amdur, M.O., Doull, J., and Klassen, C.D., 1991. Casarett and Doull's Toxicology, 

4th Edition, Pergamon Press, New York, pp. 47-48. 

25. Osweiler, G.D., Carson, T.C., Buck, W.B. and Van Gelder, G.A., 1985. Clinical and 

Diagnostic Veterinary Toxicology, Kendall/Hunt Publishing Co, Dubuque, IA, pp. 

451-453. 

26. Amdur, M.O., Doull, J., and Klassen, C.D., 1991, Casarett and Doull's Toxicology, 

4th Edition, Pergamon Press, New York, pp. 47-48. 
27. National Marine Fisheries Service, Southeast Fisheries Center, Charleston Laboratory, 

1993. Forensic Manual (for the Analysis of Marine Animal Tissues), [Drafft], pp. 98-106. 

28. Stegeman, J. J., Hahn, M. E. and Moore, M. J., (undated) Tissue Request and Sampling 

Protocol for Freshly Dead Marine Mammals, Woods Hole Oceanographic Institution, 

Woods Hole, MA, 1 page. 

29. National Marine Fisheries Service, Southeast Fisheries Center, Charleston Laboratory, 

1993. Forensic Manual (for the Analysis of Marine Animal Tissues), [Draft], p. 14. 

30. National Marine Fisheries Service, Southeast Fisheries Center, Charleston Laboratory, 

1993. Forensic Manual (for the Analysis of Marine Animal Tissues), [Draft], PP. 74-75. 

31. Geraci, J. R and Lounsbury, V. J., 1993. Specimen and Data Collection, Marine Mammals 

Ashore: A Field Guide for Strandings, A Texas A & M Sea Grant Publication, Galveston, TX, 

p. 204. 

32. Strafuss, R.C., 1992. Necropsy, in Laboratory Procedures for Veterin~y Technicians, P.W. 

Pratt, (ed.), American Veterinary Publications, Inc., Goleta, CA, pp. 471-474. 

33. U.S. Department of Agriculture, Animal and Plant Health Inspection Service, Veterinary 

Services, 1992. Directory of Animal Disease Diagnostic Laboratories, in dooperation with the 

American Association of Veterinary ~aboratory Diagnosticians (AAVLD), July.