UGA Zoo and Exotic Animal Pathology Service, University of Georgia, Athens, GA, USA
Introduction
A thorough, well-documented necropsy is a valuable opportunity to answer questions about the health of the individual animal, confirm suspected health problems, indirectly learn the health status of the group/flock/herd, and validate adequacy of parasite control and other preventive medicine practices. As a former zoo clinician and now zoo/exotic animal pathologist serving small and large zoos, aviaries, mixed species collections, and private practitioners with a wide range of species, I often see the benefit of a necropsy to both clinicians and pathologists. Gross necropsy with histopathology often provides "closure" to both the client and the exotic animal clinician, regarding health status. Occasionally, unexpected problems in health or husbandry practices are revealed, which would not otherwise have been detected. Ideally, such discoveries will allow corrective actions to minimize impact on other animals in the group and improve future care. Necropsies of deceased rehabilitant or free-ranging wildlife can reveal many important environmental quality issues or emergent diseases.1 Captive management programs (e.g., Species Survival Programs [SSPs]) in accredited AZA institutions rely on volunteer veterinary clinical and pathology advisors to review necropsy reports, identify disease trends, and advise on preventive medicine and management strategies. (See the Great Ape Heart Project [GAHP] at greatapeheartproject.org for one example of how AZA and AAZV-associated clinicians and pathologists detected cross-species trends in ape cardiac disease, and are now working together to improve diagnosis, treatment and prevention.)
Timely prosection by a clinician, instead of shipping the body to a pathologist, will reduce potential negative impact of postmortem autolysis and bacterial growth. This is particularly important in small birds, amphibians, and fish. However, it shifts significant responsibility for identifying lesions, taking correct fixed samples for histopathology and fresh samples for ancillary tests, and ensuring biosecurity (control of potential pathogens, and protection of staff and other animals on site) to the clinician prosector. Proactive teamwork, consultation and planning between the prosector and the histopathologist, will reduce suboptimal handling of tissues and maximize the chance that the pathologist can answer the clinician's diagnostic questions.
Necropsy Preparation: Location, Equipment, Personnel, Resources, Protocols
Keep the body refrigerated; never freeze it. Pick a necropsy location with controlled foot traffic (even if only controlled by a sign on the door) which will minimize spread of pathogens to people and animals and allow post-necropsy cleaning and disinfection; postponing necropsies to late evenings or weekends may help. When ready, gather adequate personnel and instruct them regarding personal protective equipment (PPE). Ideally, this involves separate clothing or lab coats which can be laundered on site, gloves, masks (when appropriate to protect from airborne pathogens), plastic apron, head and foot covers or boots. Consider eye protection, due to hazards from cutting (especially power tools). Obtain equipment (Stryker saw for chelonian shells or bone, fine scissors and forceps, cutting or pin board for small animals) to access, move and manipulate the anatomical parts. Organize sufficient collection materials and containers to avoid moving in and out of the room for supplies. Arrange scrub pans, boots, brushes, tools, water and appropriate disinfectants which are active against the potential pathogens. For instance, tuberculocide will be needed for mycobacteriosis, instead of standard quaternary ammonium compounds. Potential surprises (lesions suggesting possible zoonotic disease, or later legal or client challenges) should be proactively anticipated and personnel should have access to more stringent PPE if desired, or the number of personnel should be limited.4,10 This is especially important with zoonotic disease, such as potential rabies or tuberculosis. Photodocumentation, scales for weighing, and good note-taking are essential tools. Use of a biosafety hood is advised for birds or nonhuman primates, and in any situations where aerosolization and spread is likely (particularly fungal spores or viral infections).
Anatomical guides for prosection are available on the internet and through www.exoticdvm.com. The Southeastern Cooperative Wildlife Disease Study (SCWDS) publishes an excellent field manual for identification of gross lesions in wildlife. Standardized "best practices" necropsy protocols for many SSP species are posted in member's sections of AZA and AAZV websites, and on several SSP or Taxon Advisory Group (TAG) websites and provide narrative explanations, sample worksheets, tissue checklists, and resources. A publicly accessible publication for the World Organization for Animal Health (OIE) gives an example worksheet.11 The CL Davis DVM Foundation (a nonprofit veterinary pathology organization) sells an outstanding small paperback necropsy manual for domestic animals with excellent diagrams and worksheet examples.7 Using these resources as templates, the clinician and pathologist can create species-tailored report forms and checklists for fresh and formalin tissue collection using standard word-processing programs. These protocols can be adapted or expanded for special considerations (zoonotic disease, flock health screening). The SSP standardized protocols can be long (and some are onerous), but this is because they are optimized full necropsy protocols, intended to satisfy population-wide survey needs as well as individual animal diagnosis. Practitioners may need to prioritize some samples, and such prioritization is not unusual. For example, brain and head are routinely sampled in academic and diagnostic laboratory settings, but not all institutions routinely perform a full spinal cord examination unless neurological signs (or clinician request) warrant it. The necropsy worksheet and tissue checklists can be laminated and taken into the necropsy room where dry-erase pens are used to mark them. At the end of the necropsy, these can be photographed, and the originals can be cleaned, disinfected and re-used. Many of our clients incorporate wording or check-off boxes from such checklists into their medical records and attach them to our submission sheet. They identify the fixed and fresh-frozen samples which were taken, and which samples were submitted (including any "unknown tissues" or gonads). They note which tissues are retained on site for possible ancillary tests, and which are sent to us or other laboratories for culture, serology, or other tests. Leaving a space for special questions that the clinician wants the pathologist to answer (such as gender determination) is especially appreciated.
Performing the Necropsy
It does not matter whether the animal is a companion bird, small mammal, reptile, amphibian, fish or elephant: a necropsy (just like an antemortem physical examination) should be consistent and thorough and ensure that all body systems are evaluated and sampled, consistent with diagnostic needs. The necropsy leader/prosector should review the animal's history, situation and symptoms; formulate a list of diagnostic questions and potential differentials; plan which samples are needed a) for baseline information, b) to confirm or refute those differential diagnoses, and c) to be comprehensive enough to find unexpected problems. Be ready to explore other tangential possibilities that the necropsy may reveal, without being distracted from your overall goal(s).
Gross Necropsy Examination (Based on Small Mammals)
The prosector acts as the eyes, ears, hands (and nose!) of the histopathologist. Photos and reports of gross necropsy findings maximize the chance that gross and histopathology results can be correlated together, for accurate diagnosis. Photos will help orient the pathologist to better imagine the entire lesion, and its impact on surrounding living tissues, before that piece of tissue hardened into a brown hard lump, from which only a single 0.5-micron slice appears on a slide. Some clients use smartphones or tablets which are encased in protective layers (for biosafety) which can be disinfected. Some use disposable or waterproof/disinfectable cameras from which the memory card can be extracted. Always include a small ruler (but not on the tissue itself!) on the edge of the picture, dark background if possible, and orient the camera parallel to the tissue, approximately 6 inches away. A good well-focused picture from farther away (it can be digitally enlarged if adequate megapixels are used), is better than a blurry close-up.
A thorough external examination should note leg bands, other markings or lesions. Take body weight and judge musculoskeletal condition, degree of autolysis and dehydration. Before opening thoracic or abdominal cavities or joints, have sampling equipment ready (slides for smears, syringes, needles, red-top and EDTA tubes, swabs, conical tubes) for collection of fluid, blood, pus or abnormal joint fluid for cytology, culture or PCR. Proceed in a consistent systematic manner through anatomical areas (external areas and superficial lymph nodes, thoracic cavity, abdominal cavity - see the protocol references above for guidance). Work from clean to dirty areas (for instance, sample the gastrointestinal [GI] tract last).
Tying off proximal and distal ends, when removing thoracic pluck or abdominal tissues, will minimize leakage. Evaluate whether tissues are normal or abnormal color, consistency, texture and smell. Note if the lung pieces sink in formalin. Identify presence or absence of normal or abnormal food in the GI tract. Observe whether there is negative pressure in the thoracic cavity (if the species has a diaphragm). If the animal is large enough, assess bile duct patency by incising the proximal duodenum longitudinally on the antimesenteric surface and look on the mucosa to locate the bile duct papilla. Gently squeeze the gall bladder and note whether bile flows into the duodenum. Palpate lung, liver, and spleen, but avoid a common mistake of overmanipulating sections which will be placed in formalin. Avoid scraping the GI mucosa. Incise along airways in lungs; take serial "breadloaf" cuts through viscera such as lung, liver, and spleen (but not brain!). Collect and submit parasites for identification. Examine the heart, especially looking for developmental abnormalities in stillbirths or neonates. Refer to the GAHP postmortem heart protocol (posted on the American Association of Zoo Veterinarians website member link to necropsy protocols) for recent recommendations which modify traditional veterinary heart sampling and improve evaluation of the myocardium through transverse mid-ventricular slicing. These modifications work for any mammal, but they do not apply to avian, reptile, amphibian or fish species. Remove and freeze a small portion of brain (assuming that testing for rabies is not required) and then place the remainder intact in formalin.
Gross Exam Tips for Avian, Reptilian, Amphibian, Fish Species
Record body weight and estimate condition. For avian species, take swabs of body orifices before they are open. We routinely take a single swab of bilateral conjunctivae plus choana, then a swab of the cloaca, to save for potential PCR test for Chlamydia sp. We save a swab of the cut surface of the liver, which can be used for PCR test for viruses or Chlamydia sp. PCR. For venomous reptiles, safely remove the head into a non-compressible container (urine collection cup) before proceeding with the necropsy. For amphibians, swab skin and also swab the coelom. For fish, measure the length and girth. Prepare a fresh smear of a gill and/or fin clip and examine during or after necropsy; parasites (such as monogeneans) easily detach after death and are lost from histopathology. In some cases (particularly small amphibians or fish under 2 inches long) it is best to incise the ventral midline, save a few key fresh tissues and coelomic swabs (see following) and place the remainder of the whole body intact into formalin for decalcification and histopathology.
Selection of Samples for Ancillary Testing (Smears, Swabs, and Frozen Samples; All Species)
Diagnosis of some diseases often requires ancillary testing beyond routine H&E histopathology slides. Saving some swabs and fresh-frozen tissues as a precaution should be a priority, even before samples are taken for fixation. Having the correct archived sample for ancillary testing may determine whether an agent or differential diagnosis can be confirmed or ruled out. For example, Encephalitozoon cuniculi can be involved in cataract and lens-induced uveitis in rabbits,5 but histopathology is only rarely sufficient to detect the organism; PCR and immunohistochemistry (IHC) are often needed8. Similarly, a sample of frozen kidney is more desirable than brain, if lymphocytic choriomeningitis virus (LCMV) infection is suspected in a rodent. A PCR test of a swab of the oral cavity of a rat is the best way to document the presence or absence of Streptobacillus moniliformis (due to suspected rat bite fever in a human). Culture might be assumed to be preferable, but it is not advised due to the fastidious requirements of the organism.2 Identifying some zoonotic or herd-health-impacting diseases may require bacterial culture and serotyping.15 Immunohistochemistry may be needed for some infectious diseases, but if the antibody in the test is derived from the same species (especially mouse, guinea pig, rabbit) there may be interference.3,16 Bone marrow is useful to show whether myelosuppression or myeloproliferative disease is present in any species. Benzimidazole toxicosis in rabbits results in bone marrow suppression.6 If anemia from this or other disease is suspected, a core bone marrow biopsy must be supplemented by the hemogram (including hematocrit and morphology of the erythrocytes) and fresh bone marrow smears for cytology.12 Sugar gliders can have Chagas' disease (infection with Trypanosoma cruzi), an emergent disease problem in some sections of the United States. This may require other tests for confirmation.14
At minimum, saving fresh-frozen samples of brain, lung (or gill), liver, kidney, and intestine (tied off at either end) in separate sterile plastic bags, for any species, is advised. In tiny animals, coelomic swabs may substitute for these samples. Make fresh smears of samples for cytology; microcentrifuge tubes are useful for samples with few cells. When fluid samples are taken in syringes, remove the needle and replace with a cap, to reduce hazard to laboratory personnel. Place samples with known or suspected biohazard potential in labelled biohazard bags. When toxic agents are suspected, the best samples for assay depend on the toxin, and its site of storage or metabolism. Different assays require whole blood, GI contents, brain, fat, liver, kidney, or muscle. In small animals, limitations of small sample size may limit definitive findings. Some toxicology assays, such as at Michigan State University Diagnostic Center for Population Animal Health, may be able to use specially designed filter paper kits for dried blood spots.
Selection of Samples for Histopathology (Formalin Fixation; All Species)
Formalin penetrates very slowly. Meanwhile, autolysis is proceeding. As with fish or other small whole body specimens, ensure that formalin is bathing all aspects of the tissue, or tissues may arrive at the histopathologist unfixed and autolyzed. Cut sections of representative tissues (brain, lung, heart, liver, kidney, gonad, each division of upper and lower gastrointestinal tract, skin, muscle, and bone as a minimum). Ensure that each is no more than 0.5 inch thick. Don't let pieces of tissue stick to the sides or top of the formalin bucket (a paper towel on top will help the latter). Tissue cassettes can be used and labelled, for small tissues or masses (use pencil or special pens with formalin-resistant ink). Manila paper laundry tags can be used for labels; some prosectors are successful with paper towel or index cards; sutures can mark margins. Having enough formalin (10 parts neutral buffered formalin to 1 part tissue) is critical. Use a wide-topped non-breakable jar or bucket when possible. Incise any tissues which have thick fibrous or muscular capsules (tumors, avian ventriculus or large spleens) and open the cranium on avian heads (if placed intact in formalin) to ensure formalin penetration. Fixed samples which are often useful, but often not included, are eyes, esophagus, skin, muscle, and bone marrow in all species; nasal turbinates in rodents; osseous bullae in guinea pigs, rabbits and chinchillas; and heads of chelonians, lizards and snakes.9,13 The latter samples will require decalcification and extra time, but often show oral lesions such as stomatitis or necrosis, or bone marrow lesions which would otherwise not be seen.
After the Necropsy: Submission (and Storage) of Samples
Submission of Samples
The US Postal Service and other carriers have clear guidelines for shipping biological samples, including fixed or frozen tissues and swabs; refer to their websites for specific requirements. Use at least three leakproof levels (the primary container, enclosed in a secondary container, ideally within an intact Styrofoam container, which in turn (for some carriers) must be within an outer sealed cardboard box. Place submission paperwork in a separate closed Ziploc bag. When time is not critical for fixed specimens, it is best to allow tissues to fix well, then reduce the amount of formalin and substitute formalin-soaked paper towel, in Ziploc bag or container. During extreme hot or cold weather, ship overnight if possible, even for fixed samples. Cold packs may be needed to avoid cooking tissues in airline warehouses in case of delays; small warm packs and insulation may be needed to keep formalin-suspended tissues from forming ice crystals. Shipment of frozen tissue samples and swabs on ice packs are even more critical. If possible, avoid shipping to arrive on Fridays or weekends; unexpected travel delays could ruin your samples.
Please provide a complete history, and especially alert the pathologist to suspected differential diagnoses, or questions which the clinician wants to be answered. Always pre-warn the laboratory, and clearly mark and package any fresh samples in which zoonotic biohazard is suspected (examples: lymphocytic choriomeningitis virus, Cryptococcus sp., tularemia [Francisella tularensis], plague [Yersinia pestis]). Use leakproof, multilayered containers (see above) and clear biohazard labelling.
Storage space is at a premium everywhere; this includes academic institutions as well as practice facilities. Indefinite storage of formalin-fixed wet tissues, frozen tissues, and even paraffin blocks should not be assumed without asking. If zoonotic or legal concerns are anticipated, or publication of findings is possible, please alert the histopathologist. Proactive discussion between clinician and pathologist will ensure that everyone's needs are anticipated and met, and avoid surprises such as tissues being discarded. At minimum, tissues should be retained until the report is finalized. But some clients do not raise the possibility of zoonotic concern until after the final necropsy report is issued, because they may erroneously assume that all diseases are being screened for. Publication of cases or retrospective studies may require additional slides or tests from paraffin blocks or original tissues. Current standards in the human medical community indicate that 10 years is the required length of retention of medical records and biomaterials. Similar pressures for space in the veterinary community should be anticipated.
Acknowledgments
The author gratefully acknowledges the help of Drs. Drury R. Reavill in the preparation of this manuscript. She also acknowledges the support and assistance of Drs. Elizabeth W. Howerth, Nicole Gottdenker, and Angela E. Ellis, the Infectious Diseases Laboratory and the Department of Pathology for work which supports this manuscript and presentation.
References
1. Cruz FND, Giannitti F, Li L, Woods LW, Del Valle L, Delwart E, Pesavento PA. Novel polyomavirus associated with brain tumors in free-ranging raccoons, western United States. Emerg Infect Dis. 2013;19:77–84.
2. Elliott SP. Rat bite fever and Streptobacillus moniliformis. Clin Microbiol Rev. 2007;20:13–22.
3. Eyzaguirre EJ, Walker DH, Zaki SR. Immunohistology of infectious diseases. In: Dabbs DJ, ed. Diagnostic Immunohistochemistry. 2nd ed. Philadelphia, PA: Elsevier; 2006: 43–64.
4. Fairfax RE. Mycobacterium tuberculosis risk for elephant handlers and veterinarians. Appl Occup Environ Hyg. 2001;16:350–353.
5. Giordano C, Weight A, Vercelli A, Rondena M, Grilli G, Giudice C. Immunohistochemical identification of Encephalitozoon cuniculi in phacoclastic uveitis in four rabbits. Vet Ophthalmol. 2005;8:271–275.
6. Graham JE, Garner MM, Reavill DR. Benzimidazole toxicosis in rabbits: 13 cases (2003 to 2011). J Exot Pet Med. 2014;23:188–195.
7. King JM, Dodd DC, Roth L. The Necropsy Book. 5th ed. Gurnee, IL: Charles Louis Davis DVM Foundation; 2006.
8. Leipig M, Matiasek K, Rinder H. Value of histopathology, immunohistochemistry, and real-time polymerase chain reaction in the confirmatory diagnosis of Encephalitozoon cuniculi infection in rabbits. J Vet Diagn Invest. 2013;25:16–26.
9. McManamon R, Reavill DR. Challenges and solutions in diagnostic pathology of exotic small mammals. In: Proceedings from the annual conference of the American College of Veterinary Pathologists; October 21, 2015.
10. McManamon R, Terrell SP. Elephant postmortem examination: practical application of general principles. Invited presentation at the USDA Tuberculosis in Elephants Seminar, Kansas City, Missouri, April 5–6, 2011. Available from www.aphis.usda.gov/animal_welfare/pg.php?pg=Tuberculosis_in_Elephants (VIN editor: link could not be accessed on 12-2-15; an alternative link is www.aphis.usda.gov/animal_welfare/downloads/elephant/Postmortem%20Exam%20Procedures.pdf).
11. Mikota SK, Aguilar RA. Management protocols for animals in captive propagation and reintroduction programmes. Rev Sci Tech (OIE). 1996;15:191–208.
12. Ness RD. Clinical pathology and sample collection in small exotic mammals. Vet Clin North Am Exot Anim Pract. 1999;2:591–620.
13. Pfent CM, Mansell J, Pool RR, Mitchell ME. Pathology in practice. Disseminated idiopathic myofasciitis. J Am Vet Med Assoc. 2013;242:43–45.
14. Reavill DR. Unusual and unexpected infections: the value of additional testing. Presentation at the Association of Exotic Mammal Veterinarians Conference, Indianapolis, Indiana, September 14–19, 2013.
15. Swanson SJ, Snider C, Braden CR, Boxrud D, Wunschmann A, Rudroff JA, Lockett J, Smith KE. Multidrug-resistant Salmonella enterica serotype Typhimurium associated with pet rodents. N Engl J Med. 2007;356:21–28.
16. Taylor CR, Shi SR, Barr NJ, Wu N. Techniques of immunohistochemistry: principles, pitfalls and standardization. In: Dabbs DJ, ed. Diagnostic Immunohistochemistry. 2nd ed. Philadelphia, PA: Elsevier; 2006: 1–42.