Necropsy of Mammals

(See also Necropsy of Birds (Techniques Overview))

Video Available: Necropsy of Wild Ungulates:
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• Post mortem examination (i.e. necropsy) is an extremely valuable tool in disease investigation and management. It is important to approach each carcass with an open mind, not assuming that the cause of death is known, even if there are obvious external lesions or a known on-going disease problem.
• Before starting a necropsy, consider whether the skin or skeleton is important for museum-based studies. If it is, a cosmetic post mortem is required. (see D257 - full text available for a description)
• Reasons for performing a post mortem examination include "finding the cause of death, confirming a diagnosis, investigating unsuccessful therapy, increasing knowledge" (D91.4.w1) and for the detection of sub-clinical disease.
  • Post mortem examination is particularly important for animals which die in quarantine in preparation for introduction to a collection, translocation or reintroduction program. 
• Post mortem examinations may be performed in the field or laboratory; by the case clinician or by a specialist pathologist, dependent on circumstances.
  • Where the gross post mortem is performed by an individual other than the pathologist who will perform the further examinations on samples provided, communication between the two parties is essential to ensure that optimal samples are taken (tissue type, volume/ weight, storage, transport, temperature etc.).
  • Inadequate or incorrect sample taking may reduce the likelihood of reaching an accurate diagnosis.
  • Forensic post mortems for legal investigation should be performed by an experienced wildlife pathologist, since the credentials of the pathologist will be assessed as part of the case. (J1.32.w7)
• Similar protocols should be used for the post-mortem examination of domestic, free-ranging or captive wild mammals.
• Post-mortem examination should be conducted in good daylight whenever possible. (D286.2.w2)
• If possible, findings should be dictated during the examination, or, as an alternative, noted down at the time of the examination. (D286.2.w2)
• When performing a necropsy or post mortem examination, it is important to:

• first consider the history of the animal (where available). Note the reported clinical signs, treatment, diagnostic tests, possible differential diagnoses, number of animals involved, etc. Communication between the pathologist and the case clinician, where available, is recommended.
• consider recent and historical disease problems in the collection (captive animal), region (free-ranging), in-contact domestic animal and human populations. 
• examine the site where the carcass was found if possible (e.g. evidence of agonal movements, convulsions disturbing the local area; piles of faeces and urine around the hindquarters suggestive of prolonged recumbency).
• have a systematic approach, whether head-to-tail, system by system (digestive system, respiratory system etc.), or any other.
• recognize the normal anatomy, normal appearance of organs/tissues and anatomical variation between species.
• have knowledge of seasonal differences in the body condition and reproductive system which are normal for the species under examination.
• have knowledge of the expected variations between individuals of the same species dependent on whether they are captive or free-ranging. (e.g. obese body condition may be seen in captive animals but is unlikely in free-ranging individuals; ectoparasite and endoparasite burdens may be expected to be greater in free-ranging wild animals than those under captive management.)
• have knowledge of the method, route and time of euthanasia if performed.
• have knowledge of potential artefactual findings e.g. hypostatic congestion (pooling of blood in organs under the effects of gravity which can be mistaken for pathological congestion), barbiturate crystals from euthanasia solution which can be mistaken for gout (See: Gout in Waterfowl), pseudo-prolapse of the anus or vagina as a result of increased pressure within the abdomen caused by gas production after death.)
• accurately describe lesions/abnormalities.
• record both positive and negative findings.
• keep accurate records, including a unique identifying number for each carcass and for samples from that carcass.
• keep detailed notes on all findings and procedures for forensic post mortems, written in non-technical language wherever possible, for use in court. (J1.32.w7)
• avoid the use of non-standard abbreviations in permanent records.
• take photographs (include case identification details) for animal identification and illustration of gross pathology, particularly if the case may be involved in a prosecution enquiry. (J1.32.w7)
• preserve samples (tissues, parasites etc.) for further testing and future reference/research. 
  • A full spectrum of samples should be taken, where possible, at the initial examination if possible and stored appropriately. 
  • Further investigations, at first, may be directed at the samples thought most likely to be important in revealing the cause of death. However, if further samples are needed subsequently, the full spectrum are available in store.
  • Where time or financial constraints limit sample taking, a short list of standard tissues should be sampled, in addition to those with apparent gross pathology. 
  • In some circumstances it may be advisable to keep the entire carcass for a period following the post-mortem examination, refrigerated in the short term and frozen in the long term, to provide samples in the future if required.
• consult the appropriate regional authority if a notifiable disease (e.g. Foot-and-Mouth Disease) is suspected before progressing with the post mortem examination, 
  • Consult the List A and List B of notifiable diseases made available by the Office International des Epizooties - World Organisation for Animal Health (See: Website Ref - W31 - Office International des Epizooties)

• Carcass location and body size may dictate whether transport to the laboratory facility for examination is possible, or whether the post mortem must be performed in the field.
• Where field post mortem examination is unavoidable, attention should be paid to the risk of spread of infection to wild or domestic animals through opening of the carcass and available methods for carcass disposal (e.g. pit, cremation).
• Autolysis of the organs occurs with variable speed; the adrenal medullae, gastro-intestinal mucosa, pancreas, liver, kidney and central nervous system develop autolytic changes particularly quickly.
• Post mortem examination should be performed as quickly as possible after death has occurred and has been confirmed. However this may not always be possible, and carcass cooling to slow the rate of autolysis should be practised. 
  • Some authors suggest soaking of the fur in cold water with a small amount of detergent to aid in wetting of the skin.
  • The carcass should be placed within a sealed plastic bag, clearly labelled, with excess air removed, and be refrigerated if its body size allows.
  • Carcasses preferably should be refrigerated while awaiting examination. (B10.3.w18)
    • Where sufficiently large refrigeration facilities are unavailable, the carcass should be moved to as cool an area as is available.
  • With very large mammals, cooling of central organs will not occur sufficiently quickly to prevent autolysis; priority should be given to performing the post mortem as soon as possible; opening the abdomen may help lower the core temperature as quickly as possible.
  • Where post mortem examination must be delayed until 72-96 hours after death, the carcass should be refrigerated only. However if the examination must be delayed over 96 hours post mortem, it is recommended to freeze the carcass immediately.
• When transporting a carcass or pathological sample to a laboratory for analysis, attention should be paid to temperature control in transit. Insulated containers should be chosen, ice packing of frozen samples may be used and times when postal delays may be expected should be avoided (e.g. weekend, public holidays, strikes).
  • Local regulations governing the postage of pathological samples should be consulted (labelling, courier, container type etc.)
• In the event of a die-off (mass mortality event) it is important to examine fresh carcasses of a number of individuals, representative of the range of species affected and the ages of individuals affected, and to remember that more than one disease process may be acting at any one time and that the major cause(s) of death may change during a prolonged die-off.
• The results of the post mortem examination should be used in conjunction with the history of the mammal or mammals and assessment of the environment to help determine their significance and recommended future action. 
• In areas where rabies infection (See: Rabies) is enzootic, all mammals found dead, and particularly those with a clinical history of abnormal behaviour or neurological signs, should be carefully examined and considered as potentially infected until proven otherwise. (See: Human Health Considerations section on this page)
• Suspect cases of sudden death should have peripheral blood smears taken to exclude anthrax infection (See: Anthrax) as a differential before the carcass is opened. Bloody discharges should direct the examiners' attention to the need to exclude anthrax infection before continuing with the examination. Dependent on region, specialist veterinary staff may be legally required to carry out the anthrax testing process.
  • Samples should be taken by nicking the dependent ear or from the coronary band. 
  • In wild equids (Equidae - Horses (Family) - horses and zebras), wild pigs (Suidae - Pigs (Family)) and carnivores (Carnivora - Carnivores (Order)), anthrax bacilli may not be present within the blood, therefore examination of a smear made from the cut surface of a lymph node (usually submandibular) is recommended in addition.
  • Tissue and blood smears should first be air dried and then be fixed in methanol.
  • Staining should be performed for two minutes with polychrome methylene blue, or Giemsa stain.
  • Samples should be examined under oil immersion microscopy for evidence of anthrax bacilli.
  • If anthrax infection is confirmed, careful attention must be paid to quick and effective carcass disposal. Regional authorities responsible for disease control should be notified and action taken as appropriate.
  • If anthrax infection is excluded, the post mortem examination should proceed.

  (B411)

• Estimation of time of death is not as widely a developed skill in wildlife as with human pathology; forensic entomology has not been used extensively in wildlife cases to date. (J1.32.w7)

• Detailed knowledge of ballistics (shot gun, air gun, arrow) and the typical wounds that they cause is useful, particularly for forensic post mortem examination. (J1.32.w7)

• Knowledge of the species of common predators for the animal under post mortem examination in that region is useful. An understanding of the distribution of the wounds that they typically inflict can be a useful aid for identification of cause of death or scavenging. (J1.32.w7)

• Note: If poisoning (e.g. plant poisoning) is suspected, the pathologist should be informed of this suspicion before the necropsy is carried out. (P6.4.w1)

• For information on carrying out a Cosmetic Post Mortem, to enable the skin and skeleton to be used for museum-based studies see: Cosmetic Post Mortem

(B10.3.w18, B127, B273, B411, B433, J1.32.w7, P24.327.w13, D91.4.w1, D257 - full text available, V.w26)

• Post mortem examinations may be performed on hedgehogs which have been found dead in the wild, those which have died shortly after presentation as a casualty, those which have died after treatment and a significant period in care, and those which are found dead shortly after release.
  • Consideration should be made as to whether the hedgehog has a recent history of free-ranging or captive management.
  • Certain conditions are more likely to develop after a period in captivity e.g. obesity, possible dental disease (See: Hedgehog Dental Disease)
• Use the season, age and sex of the hedgehog to guide investigation to the most probable causes of death.
  • e.g. starvation or hypothermia during the winter. (See: Starvation), burns during November close to Guy Fawkes night (bonfire night) in the UK. (See: Burns and Smoke Inhalation)
• When found dead in the wild, use study of the local habitat and hazards to suggest possible causes of death. e.g. roads, ponds, cattle grids. (See: Garden Management for Hedgehogs (Erinaceus europaeus))
• Ensure knowledge of the normal anatomy of the hedgehog. (See: Erinaceus europaeus - West European Hedgehog) 
• Knowledge of the marked variation in hedgehog body systems with season is required:
  • Seasonal variation in body weight, condition and fat deposits; particularly before and after hibernation. (See: West European hedgehog - Appearance-Morphology- Measurement and Weight, )
  • Seasonal variation in the male reproductive system with significant growth in the accessory sex glands in the breeding season with regression to inactivity in the autumn. (See: West European hedgehog - Detailed Anatomy Notes (Literature Reports) and West European hedgehog - Life Stages (Literature Reports) - Male Seasonal Variation)
• Hedgehogs are susceptible to natural infection with Foot-and-Mouth Disease, a notifiable disease. Where infection is suspected, the appropriate regional authorities must be contacted. (See: Foot-and-Mouth Disease)
• Arranging a hedgehog into a spread-eagled position immediately after death (i.e. before rigor mortis sets in) may be useful if a post mortem examination is to be carried out. (B337.6.w6)

• Necropsy of an elephant "is a laborious task." (D286.Intro.w9)
• The constraints associated to the medical management of diseases in elephants are also applicable to the post mortem examination. (J359.7.w2)
• The opportunity to perform the post mortem examination before the autolytic changes begin is rare, particularly in free-ranging elephants. (J359.7.w2)
• If necropsy is to be delayed, consider covering the carcass with ice. (D292)
• Elephant necropsy should not be attempted by a single person; at least two assistants are required (i.e. a minimum team of three people). (P80.1.w1)
• The autopsy should be conducted systematically by a team of experts, ideally a pathologist, microbiologist and parasitologist along with skilled personnel to assist in the cutting and opening of the body, with the objective of making a specific disease diagnosis. (J359.7.w2)
• Assigning specific tasks to each of the necropsy team members is recommended. (B450.B.w28, D292)
  • It may be useful to designate a media liaison to act as spokesperson for enquiries from the media. (D292)
• Access to a clinical history and medical management of the elephant is a very valuable tool to achieve the post mortem objective of making a specific disease diagnosis. (J359.7.w2)
• Always consider the animal's past clinical history and treatments given, as well as the recent signs from physical examination, diagnosis made, treatment and the manner of its death. (D286.2.w2)
• Necropsy should always be attempted; it is acknowledged that it may not be possible to gather much useful information if the carcass is already decomposed. Decomposition starts quickly, particularly in hot weather. (D286.Intro.w9)
• In the field, approach a "dead" elephant which is in lateral recumbency carefully: it may be asleep and wake and attack when approached. If vultures are present, it is probably dead. (B411.IV.w4, P80.1.w1)

ALWAYS CONSIDER:

• Anthrax: DO NOT conduct a post mortem examination if anthrax is suspected. (B101, B336.67.w67, J64.21.w21)
  • Prior to starting the examination, make a blood smear and examine it to check for anthrax. (D286.2.w2)
• Mammalian Tuberculosis.
  • Search intensely for TB lesions during any elephant necropsy, even if the circumstances of death or euthanasia do not give any reason to suspect TB. (D292 - FULL TEXT INCLUDED)
  • Open the thoracic cavity last; this should be carried out by personnel wearing appropriate personal protective equipment such as hepa-filter masks. (D292)
• Herpesvirus Infection in Elephants For information on taking and sending samples, see:
  • D289 - Instructions for Elephant Endotheliotropic Herpesvirus (EEHV) Sample Submission - FULL TEXT INCLUDED

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D293 - Elephant Research And Tissue Request Protocol

• Autolysis is rapid, particularly of the abdominal contents. Necropsy should take place as soon as possible, preferably within four hours of death. (B600.17.w17, B611.4.w4, B614.5.w5, J72.49.w4)
  • This is particularly important if investigating suspected intestinal disease. (B600.17.w17)
  • Autolysis can be slowed by refrigeration. (B600.17.w17)
• As with necropsy of other species, a systematic approach is important; a preprinted necropsy checklist can be advantageous. (B600.17.w17, B611.4.w4)
• Samples should be collected for histopathology (often needed for confirmation of diagnosis) and/or bacteriology. (B600.17.w17, B611.4.w4) Samples of the following tissues should be taken:
  • Heart (the whole heart can be submitted).
  • Liver.
  • Kidney.
  • Spleen.
  • Lung.
  • Lymph node.
  • Intestines.
  • Note: It may be useful to provide sections of both grossly normal and visibly abnormal sections of organs (e.g. liver). (B600.17.w17)
• Sections chosen for examination should be representative of the lesion and preferably should also contain some adjacent tissue which appears grossly normal. (B611.4.w4)
• Cut sections 6 - 10 mm thick to ensure adequate penetration of fixation solution. (B611.4.w4, B614.5.w5)
• Usually, 10% neutral buffered formalin (formal saline) is used for tissue preservation. (B611.4.w4, B614.5.w5)
• Note: investigations of disease in wild burrowing rabbits such as Oryctolagus cuniculus - European rabbit are complicated by the fact that sick rabbits commonly die in their burrows, while those which do die above ground are rapidly removed by predators. This reduces access to carcasses for necropsy. (J1.30.w9, J1.34.w13, P2.47.w4)

• Post mortem examination may provide valuable information, particularly in the event of an outbreak of disease. (B232.16.w16)

All great apes in a collection which die should be necropsied as soon as possible. (D428.8.2.w8b). Because of the small number of bonobos in captivity and the relative lack of information on their diseases, it is important to gain all possible information from any bonobo necropsy. A protocol for great ape necropsy should be followed, such as the AAZV "Standardized necropsy report for great apes and other primates" (D408)

The following information is appropriate for all of the great apes:

• Note: 
  • Standard sections should be taken and fixed from normal organs as indicated in the sections following. Additionally, sections of all lesions should be preserved.
  • If possible, two sets of sections should be taken, one to be sent for histopathological examination for diagnosis, and the other kept for archiving (in the USA, by the SSP pathologist). (D408)
  • Specimens should be labelled. For specimens which may not be identifieable (e.g. lymph nodes, either place each into a labelled embedding bag, or cassette, or leave attached to identifiable tissue. (D408)
  • Hollow viscera and skin can be allowed to attach flat for a moment to (labeled) pieces of paper (serosal surface touching the paper) before being placed, with the paper, into the formalin. (D408)
  • Label the container with the identity of the animal, age, sex, date, location and the name of the person carrying out the post mortem examination. (D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Laboratory post mortem facilities should be housed in a separate room to the clinical facilities and should be easily cleaned, with adequate water supply and draining floors. 
  • Requirements for air exchange systems, fume cupboards etc. will vary according to the taxa under examination and, as such, the pathogen classes which are likely to be encountered.
• Suitable preventive clothing (See: Human Health Considerations section on this page)
• Weighing scales and measuring devices (tape, calipers) should be available with a level of accuracy and range appropriate for the mammal under investigation.
• Suitable dissection instruments should be available in the laboratory or the field:
  • Two suggested equipment lists for post mortem examination in the field include: 1) a curved knife for skinning; a straight, pointed knife for dissecting; a pair each of 25 cm rat-toothed forceps, 15 cm pointed forceps and 15 cm dissecting scissors; a sterile scalpel and blades; an enterotome; a bone saw; a large pair of bone forceps or bone-cutting shears; an axe; a sharpening stone and steel; a spring balance to weight up to 10 kg; a block and tackle; some nylon rope; and a small gas or alcohol burner for sterilising instruments. (B411.I.w1); 2) "knife, sharpening steel, forceps, scalpel and blades, small (hack) saw, disposable rubber gloves, rubber apron, small butane torch or alcohol lamp, matches, sterile syringes and needles, sterile swabs with transport media." (B127)
  • Dissection equipment should be clearly marked and kept solely for the purpose.
  • Equipment should be cleaned and sterilised following use.
  • Size of equipment should be tailored to the carcass size; ophthalmology instruments, hand lens and dissecting microscopes may be useful for small mammals where available.
  • All non-disposable cutting instruments should be kept sharp.
• Suitable sample collection equipment and disposables should be available in the laboratory or the field:
  • An equipment list for specimen collection in the field includes: sterile disposable 5 ml syringes and sterile needles (20 gauge); culture tubes with sterile swabs; microscope slides in box; sterile Universal bottles; sterile blood tubes; plastic bags with closure tops (Whirlpack or Ziploc type); heavy duty plastic sealing tape; 300 mL wide mouthed glass and plastic jars; a measuring tape or ruler rubber or plastic gloves; aluminium foil; a rabies kit (World Health Organisation (WHO)) (or a drinking straw in a small jar of buffered glycerine); labels, string and a waterproof marker pen or pencil. (B411.I.w1)
  • Microscope, clean slides and cover slips.
  • Sterile swabs and transport media.
  • Gas source for heating metal blade to sear surface of an organ before sampling for microbiology.
• Suitable fixation medium should be available in the laboratory or the field including 10% neutral buffered formalin, 70% alcohol for parasites, 100% acetone for cytology (danger flammable!), normal saline.
• Stain kits for cytological, bacteriological and fungal examinations (Gram, Ziehl-Nielsen, Diff-Quick^®, Hemacolor^®, Lactophenol blue, etc.). 

(B273, B411.I.w1, P24.327.w13, D91.4.w1, V.w26)

• Small dissecting instruments may be useful, particularly for post mortem examination of hoglets.
• Accurate electronic scales and calipers are required for biometric and weighing measurements.

• To move, manipulate or transport a elephant carcass a mechanical crane or a tow truck and chains may been used. (B450.B.w28, J359.7.w2)
• Chains and a tow truck may be sufficient for movement a short distance; (D292)
• A truck with a hoist is required to move the carcass to a remote location (placing it on a flatbed trailer may make this task easier). (D292)
  • If moving on a flatbed trailer, strap the elephant to the trailer and cover with a tarpaulin. (D292)
• It is important to check that the vehicle will handle the elephant's weight. (B450.B.w28, D292)
  • (Allow 2,300 - 3,700 kg for a female Elephas maximus - Asian Elephant, 3,700 - 4,500 kg for a male Elephas maximus - Asian Elephant, 2,300 - 4,000 for a female Loxodonta africana - African Elephant, 4,100 - 5,000 for a male Loxodonta africana - African Elephant. (D292)
  • Information on normal body weights in elephants is provided in:
    • Elephas maximus - Asian Elephant
      • See: Asian Elephant Elephas maximus - Appearance-Morphology- Measurement and Weight (Literature Reports)
    • Loxodonta africana - African Elephant
      • See: African Elephant Loxodonta africana - Appearance-Morphology- Measurement and Weight (Literature Reports)
    • Loxodonta cyclotis - Forest Elephant
      • See: Forest Elephant Loxodonta cyclotis - Appearance-Morphology- Measurement and Weight (Literature Reports)
• If the transportation of the carcass is delayed, the animal body should be covered with ice. (B450.B.w28, D292)
• A recommended equipment check list includes: (B450.B.w28)
  • Large animal necropsy instruments, including large knives, scalpel handles and scalpel blades, forceps, iron spatula, surgical scissors. 
  • Retractors of different sizes and shapes.
  • Sterile instrument for culture samples.
  • 10% neutral buffered formalin.
  • 4% glutaraldehyde.
  • Containers for sample collection.
  • Culture swabs and universal pots for urine.
  • Serum tubes for blood and urine collection.
  • Aluminium foil and plastic bags for freezing tissue samples.
  • Labels and waterproof marker pens.
  • Scales.
  • Tape measure, at least 2m long.
  • Chainsaw and axe (to cut through the skull); both a large and a small axe. 
  • Hammers, chisels, hand saws and hack-saws.
  • Hoist and/or crane.
  • Ropes.
  • Crow-bar.
  • Carts on rollers to move body parts.
  • Coveralls, boots, gloves, caps, protective eye and head gear.
  • Surgical masks approved for TB exposure, for all personnel involved in the necropsy.
  • Access to water with a hose.
  • First aid kit.
  • Camera and appropriate accessories (e.g. film, batteries.)
  • Emergency lamps/generator; torches (flashlights). (D286.2.w2)

  (B450.B.w28, D286.2.w2, D292)

• In the field, recommended equipment includes: (B411.IV.w4)

  "1. Four or five large butcher knives and sharpening steel or other sharpening device
  2. Several robust meat hooks
  3. A large saw, suitable for sectioning some of the massive bones
  4. A large axe
  5. A shovel
  7. A block and tackle where possible
  8. A wheelbarrow
  9. Adequate water
  10.Large PVC tray/containers and cutting boards
  11.Thick ropes or chains
  12.A portable gantry, where possible
  13.A metal detector to locate bullets in poached animals."

• Where a gantry and block and tackle are not available, a 4x4 drive vehicle with ropes and a tree fork to use as an elevation point may be able to substitute. (B411.IV.w4)
• Note: A chain saw is useful, but tends to get blocked up with sinews and fat. (B411.IV.w4)
• Camera: particularly if litigation is a possibility. (B411.IV.w4)
• Magnifying glass. (D286.2.w2)
• Torch (flashlight). (D286.2.w2)

Another list includes knives and sharpening steel, hooks, both saw and chain saw, an axe, a shovel, block and tackle, a wheel barrow, rope or chain and an adequate supply of water. (P80.1.w1)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Necropsy knife.
• Scalpel and scalpel blades.
• Toothed and rat-toothed forceps.
• Blunt-blunt, blunt-sharp and heavy (necropsy) scissors.
• Rib cutters.
• Bone saw.
• 1 mm diameter probe.
• Metric ruler.
• 10 mL syringe with needle.
• Scales (balance) on which whole animal and organ weights can be recorded.
• String (for tying off vessels and organs).
• Containers with appropriate fixative (10% formalin).
• Swabs for bacteriological samples.
• Sterile glass containers (for collection of fluids).
• Glass slides (to make impression smears from organs)

(B611.4.w4, B614.5.w5, V.w5)

Note: it is useful to have an assistant present both to help with dissection and to record findings. (B611.4.w4, B614.5.w5)

• A cork board is useful for pinning out the carcass. (B232.16.w16)
• A set of instruments should be kept for necropsy. (B232.16.w16)

Pan paniscus - Bonobo and Pan troglodytes - Chimpanzee are the closest living relatives to humans. The risk of zoonotic disease must be considered and appropriate containment facilities and personal protective equipment used.

The following information is appropriate for all of the great apes:

• Necropsy of primates should be carried out in a room which has been designated for necropsies. (D410)
• The floor and the work surface should be impermeable and easily cleaned. (D410)
• Preferably the necropsy should be carried out in a biological safety cabinet with laminar air flow. (D410)
• Appropriate personal protective clothing should be available for (and worn by) all personnel involved in the necropsy. This includes: (D410)
  • Waterproof boots or boot/shoe covers. (D410)
  • Double gloves. (D410)
    • Also an anti-cut glove on the non-cutting hand. (V.w150)
  • Coveralls or surgical gown made from a material impermeable to water and blood (e.g. Tyvek) (D410)
  • Face mask or respirator. (D410)
    • When conducting a necropsy on a primate which is too large to fit into a safety cabinet, it is recommended that a powered respirator with full face visor and hood should be worn. (V.w150)
  • Face shield. (D410)
  • Hair covering (and beard covering). (D410)
• Observers also should wear appropriate PPE including boot/shoe covers, masks, eye protection, lab coat or similar. (D410)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Record the sex and estimate age. Classification as a neonate/ infant/ juvenile/ adult/ geriatric may be useful.
• Record any identifying numbers e.g. tattoo, tags, microchip numbers etc. Retain any physical markers for future reference e.g. radio-transmitters, tags.
• Record any characteristic features e.g. characteristic scars or colour marks, coat colour (e.g. albino, leucistic, melanistic).
• Examine the inside of the container/wrappings that the carcass has been presented in, noting contaminants (e.g. mud, oil) and possible external parasites such as fleas, lice or mites which may have left the host. Presence of maggots within the wrappings may indicate significant carcass decomposition but should alert the examiner to search for lesions of fly strike (See: Myiasis)
• Weigh the carcass . Dry weight should be taken where possible to minimise error. Where the carcass is presented wet, this should be noted for future reference to allow recognition of potential bias.
• Biometric measurements: A range of biometric measurements should be taken using graduated calipers. The accuracy and units of measurement should be clearly noted. Examples include tibial length, crown-rump length. Description of the exact biometric parameter used should be available to avoid confusion in the future when trying to reproduce the measurement and compare between individuals of the same species.
• Body condition - use a combination of subjective and objective measurements of body condition where possible. 
  • Subjective scoring systems can be developed using indices based on muscle bulk, amount of subcutaneous and visceral fat deposits. 
  • Objective scores can be developed, perhaps by creating indices of parametric measurement to body mass.
• Radiography may be indicated to detect e.g. fractures, radio dense foreign bodies, air gun or shot gun pellets, metabolic bone disease.
• Examine the external surface of the carcass . Note whether the carcass is fresh or decomposed; whether it has been refrigerated or frozen; also whether it is intact or scavenged, and if scavenged to what degree. 
  • Estimate time of death where unknown.
  • Autolysis will be accelerated where the temperature of the animal was increased at the time of death e.g. heat stroke, lightning strike.
  • Differentiate gas production in the gastro-intestinal tract post mortem with pre mortem 'bloat'.
  • Bile staining of tissues adjacent to the gall bladder can be seen in carcasses as post mortem change.

(B411.II.w2, V.w26, P24.327.w13)

Sexing hedgehogs

• In the adult male, or boar, the prominent penis sheath (prepuce) is positioned in the midline close to the navel.(B262.2.w2, B260.1.w1, B228.2.w2, B254.14.w14); about five centimetres (two inches) cranial to the anus.(B254.14.w14)
• Identification of the sex of new-born hedgehogs can be challenging.(B228.2.w2) The prepuce of the male is positioned close to the anus in the infant and gradually migrates forward towards the navel as the animal grows.(B228.2.w2)
• Care should be taken not to confuse the umbilicus of the neonatal hedgehog with the prepuce of the infant male. (B228.2.w2)
• In the adult female, or sow, the anus and vagina are positioned close together (approximately 1 cm apart).(B262.2.w2, B228.2.w2, B254.14.w14)
• The testes of the hedgehog are not externally palpable or visible within a scrotum but instead are positioned inside the abdomen.(B228.2.w2, B142)
  • For further information see: West European hedgehog - Detailed Anatomy Notes (Literature Reports))

Age determination in hedgehogs

• The appearance of the spines, whether the eyes and ears are closed or open, eruption of the teeth, ability to curl into a ball, independence form the nest, etc. can be used for age estimation of infants and juvenile hedgehogs.
  • (See: West European hedgehog - Life Stages (Literature Reports) - Neonatal development)
• Body weight is too variable a parameter for use in age estimation. (B254.24.w24)
• Several techniques for age estimation have been described including examination of the growth rings in the cut surface of the jaw bone and examination of the degree of growth plate fusion.
  • For further information see: West European hedgehog - Life Stages (Literature Reports) Longevity/Mortality - Age Estimation Techniques)

Markers / Appearance

• Notes should be taken of any marking devices found on the hedgehog. e.g. radio-transmitter, shrunk on plastic markers, etc.
  • Every effort should be made to contact the organisation known, or though likely, to be undertaking a hedgehog study using such markers.
  • Feedback from marked hedgehogs may provide useful information for post-release monitoring studies, etc.
• Occasional reports of hedgehogs with unusual general appearance occur. These include true albinos, blond and spineless individuals. (B262.2.w2)
• True albino hedgehogs are seen with pink nose and feet. White hedgehogs with black nose and eyes are also observed.(B142)
• Melanistic colour variant hedgehogs with a completely black coat have never been reported.(B260.1.w1, B254.6.w6, B228.1.w1, B142)
  • For further information see: West European hedgehog - Appearance- Morphology- Skin-Coat-Pelage (Literature Reports))

Weight

• The degree of seasonal variation in body weight, condition and fat deposits; particularly before and after hibernation should be understood. 
  • (See: West European hedgehog - Appearance-Morphology- Measurement and Weight (Literature Reports))
• A scoring system based on the presence of subcutaneous fat deposits and thigh muscle bulk can be used as subjective indicators of body condition. The following scheme is based on these indicators: "1 thin, no subcutaneous fat and wasted musculature; 2 moderate, no subcutaneous fat but good muscle thickness; 3 good, some subcutaneous fat; 4 fat, a thick covering of subcutaneous fat." (J3.138.w2)

• Take pictures, particularly if litigation is a possibility. (B411.IV.w4, P80.1.w1)
• Check for signs of putrefaction. (D286.2.w2)
• Note the position of the carcass. (D286.2.w2)
  • If the positioning is awkward and a leg appears possibly broken, it is important to check that leg carefully; a fracture causing immobility can be fatal. (P80.1.w1)
• TAKE A BLOOD SMEAR BEFORE OPENING THE CARCASS, and examine it to check for anthrax. (D286.2.w2, P80.1.w1)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D293 - Elephant Research And Tissue Request Protocol

Sex and genitalia

• Identify the animal's sex. (D286.2.w2, P80.1.w1)
  • In neonates, the distance from the umbilicus to the genital orifice is about four finger-widths in males, eight in females. (B411.IV.w4)
  • The clitoris of the female is pink, the penis of the male is blueish grey. (B411.IV.w4)
• Examine the mammary glands in females, the prepuce and penis in males. (D286.2.w2)
  • See: Priapism in Elephants

Body measurements

• Body measurements can be recorded before euthanasia, if it is planned, or as soon after death as possible. (B450.B.w28, D292)
• All measurements should be taken in a straight line, unless specifically indicated otherwise. (B450.B.w28)

Weight

• Weight should be recorded when possible.

Elephas maximus - Asian Elephant

• See: Asian Elephant Elephas maximus - Appearance-Morphology- Measurement and Weight (Literature Reports) 

Loxodonta africana - African Elephant

• See: African Elephant Loxodonta africana - Appearance-Morphology- Measurement and Weight (Literature Reports)

Loxodonta cyclotis - Forest Elephant

• See: Forest Elephant Loxodonta cyclotis - Appearance-Morphology- Measurement and Weight (Literature Reports)

Body condition

• This should be assessed. (B411.IV.w4, P80.1.w1)
• Temporal depression, protrusion of the ribs or scapular spines, and a sunken flank (deep lumbar depression) may indicate poor condition. (B455.w1, B411.IV.w4, D286.2.w2, P80.1.w1)
• Very loose skin, with a "baggy pants" appearance of the skin over the hind quarters also indicates poor condition. (B411.IV.w4, D286.2.w2)
• A body score for Elephas maximus - Asian Elephant has been developed. (B455.w1)

Time of death

• This may be estimated using a combination of findings: (D286.2.w2)
  • Rigor mortis usually appears within one to four hours and lasts for 16 - 18 hours. However, it may start immediately or be delayed by four to 24 hours, and may pass off within three hours or remain for as long as 48 hours. (D286.2.w2)
  • Decomposition usually starts in six to 36 hors but is affected by the temperature and humidity. With advanced decomposition, the muscles become pale red and watery. (D286.2.w2)

Excreta and secretions

• Collect a fresh dropping, or a faecal ball from the rectum; examine for the consistency of the food particles. (D286.2.w2, P80.1.w1)

  • Coarse, poorly digested food particles in the faeces may be found in elephants with tooth problems. (B10.34.w42, B23.79A.w11, P1.1982.w1, P80.1.w1) See: Molar Maldevelopment and Malocclusion in Elephants

Presence of penetrating wounds

• These may be from bullets or tusks. (P80.1.w1)

In neonates

• Examine the umbilicus. (D286.2.w2)

Measurements

• Record the bear's weight.
• The total body length, tail length, hindfoot length (with and without claws) and ear length should be recorded.

External genitalia

Abnormalities of the external genitalia reported in bears include:

• Clitoriditis in Bears
• Cryptorchidism in Bears
• Hypospadias in a Bear
• Pseudohermaphroditism in Bears

Body condition

• Body condition in bears can be described as follows:
  • Poor condition: It is easy to feel the bear's hip bones, shoulder blades, spine and ribs; the bear looks unhealthy, like a skeleton with skin stretched over it;
  • Fair condition: The bear appears thin but not unhealthy; the bear's hip bones, shoulder blades, spine and ribs are easily palpable, but not prominent;
  • Good condition: The bear appears healthy but not extremely fat; it is difficult to feel the bear's hips, shoulder blades, spine and ribs;
  • Excellent condition: The bear appears extremely fat and healthy; the bones of the bear's hips, shoulder blades, spine and ribs cannot be palpated.

  (D249.w10)

• A five-point scoring system for polar bears (provided by the Polar Bear Specialist Group) is: (D251.5.w5)
  • 1: pelvis and scapulae protruding, ribs easily palpated, a deep hollow notable between the pelvis and last rib, showing virtually no fat.
  • 2: pelvis easily palpable, ribs palpable but with some muscle covering; an obvious hollow is present between the pelvis and the last rib, but this is softer than in (1).
  • 3: Body fully fleshed out, with obvious fat present over the pelvis and shoulders, the ribs are less obvious and there is no hollow between the pelvis and the last rib.
  • 4: the bear has a rounded or blocky appearance and is well fleshed over all bony areas, with obvious fat over the rump and shoulders.
  • 5: the bear's legs appear too short for the body; there are rolls of fat on the neck and lower shoulders.

  (D251.5.w5)

• Body weight is reduced following hibernation; some wild bears starve in the period following emergence from hibernation. See: Starvation in Waterfowl and Seabirds (with notes on Hedgehogs and Bears)

Wounds etc.

Note any localised infections or injuries:

• Abscess in Bears
• Bacterial Wound Infection and Cellulitis in Bears (Bacterial Disease Summary)
• Lacerations & Punctures, including bite wounds (with special reference to Waterfowl, UK Wildlife and Bears)
• Navel Infection in Bears
• Umbilical Hernia in Bears

• Examine the whole body visually, and palpate. (B600.17.w17, B611.4.w4, B614.5.w5)
• Consider whether radiographs should be taken before the necropsy begins. (B600.17.w17)
• Note any identifying marks or objects.
  • A ring above the hock of a domestic rabbit indicates it is a pedigree rabbit and also gives the year of birth. (B600.17.w17)

Measurements

• Record the weight. (B600.17.w17, J72.49.w4)

Sex and external genitalia

• Examine the external genitalia visually and by palpation; note any abnormalities. (B611.4.w4)

Body condition

• Note general body condition.

• Start with a general examination of the external appearance. (B232.16.w16)
• Note any discharges from orifices, any abnormalities, any indications of dehydration, diarrhoea or vomiting. (B232.16.w16)

Measurements

• Weigh the ferret. (B232.16.w16)

• Note the general condition of the carcass (e.g. fresh, decomposed) and the nutritional state of the individual.
• The external examination should look for evidence of trauma, diarrhoea and exudates.

(D410)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment
• D293 - Elephant Research And Tissue Request Protocol

• Examine the eyes, ears, nostrils (external nares) and mouth for evidence of haemorrhage, discharge, external parasites, maggots (See: Myiasis), abnormal growths, foreign bodies, other lesions etc.
  • Fatalities following acute plant poisoning may still have remnants of the plant material within their mouths.
  • Examine the eyes carefully for evidence of opacity or long standing injury which may have compromised vision. Free-ranging predators may lose condition and potentially starve if they rely on acute vision to catch prey.
• Mucous membrane colour in the mouth and eyes should be assessed for evidence of jaundice or pallor (suggestive of blood loss or profound anaemia).
• Examine the condition of the lips, oral mucosa, soft / hard palate and tongue for lesions such as ulcers, growths or developmental abnormalities (e.g. cleft palate).
• Cut through the oral commissures to allow adequate examination right to the back of the oral cavity.
• For more thorough examination, continue the incisions to allow disarticulation of the lower jaw.
  • This will enable full inspection of the nasopharyngeal area, tonsils, retropharyngeal and parotid lymph nodes, as required.
• The teeth should be examined for evidence of tooth loss, abnormal or excessive wear (attrition), gingivitis, periodontal disease, tartar accumulation etc.
  • Full dental formula should be recorded.
• Remove the skin from the skull and the temporal muscles as necessary.
  • Examine the subcutaneous tissues and skull table for evidence of trauma, bruising, etc.
• Section the base of the auricular (ear) cartilages and examine the contents for evidence of discharge, inflammation, parasites etc.
• Sectioning of the skull for examination of the sinuses, turbinates or tympanic bullae may be performed if required, given specialist facilities.

Samples

• Tissue samples, swabs for culture and impression smears should be taken when tissues are first examined and before they become contaminated. 
• Tissue to be fixed for histopathology should be no more than 10 mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Eyes may be removed with care and fixed in a preservative preferred by the histopathologist, e.g. Bouin's or Zenkers solution.
• Collect external parasites in 70% ethanol for preservation and further identification as required.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)

(B411.II.w2, B411.III.w3, P24.327.w13, D91.4.w1, V.w26)

• Closely examine the eyes, ears, mouth, and any wounds for evidence of fly eggs or maggots, particularly in the summer. (See: Myiasis)
• Check the jaw and hard palate for fractures and trauma which occur fairly frequently in hedgehogs, e.g. road traffic accident victims.
• Injuries to the nose which could have interfered with the ability to smell may be more important than damage to the eyes because the hedgehog relies principally on the senses of smell and hearing for food acquisition. (See: West European hedgehog - Feeding Behaviour (Literature Reports))
• Examine the external ear canals for evidence of mite infection. (See: Hedgehog Notoedres Mange))
• Full dental formula for the hedgehog is 3.1.3.3/2.1.2.3 (I3/2, C1/1, P3/2. M3/3).(B142)
  • For further information on dental morphology (See: West European hedgehog - Appearance-Morphology- Head and Neck (Literature Reports) - Dentition)
  • For further information on dental eruption times. (See: West European hedgehog - Life Stages (Literature Reports) - Neonatal development)
• Carefully examine the teeth for evidence of gingivitis, tooth loss, periodontal disease, tartar accumulation. These findings may be more common in animals that have been in captivity on an artificial diet for some time. (See: Hedgehog Dental Disease)

• The head is usually removed from the body and the trunk and ears are cut, to facilitate the examination. (D292, J359.7.w2)
  • The head is severed from the body at the atlanto-occipital joint. (P80.1.w1)
  • A battery-operated reciprocating saw with replacable metal cutting blade is useful for entering the cranium. Otherwise, a chain saw together with a large axe and chisels are required. (D292)
  • To remove the brain, make three connecting cuts in the flattened triangle at the base of the skull, then use a curved crow bar to remove the bony plate in pieces. Note: there is a hazard to personnel from fragments when a chain saw is used. Appropriate personal protective equipment for the eyes, face and head should be worn. (D292)
  • After the skin and muscle have been reflected, a chain saw or axe can be used to make a sagittal section just above the eyes to reach the brain. (P80.1.w1)
  • A chain saw can be used to saw the head into two pieces. (P80.1.w1)
    • This is difficult and time-consuming. (P80.1.w1)
  • A hand saw or axe can be used to open up the middle ear. (P80.1.w1)
    • This is difficult and time-consuming. (P80.1.w1)

Gross examination of the head and neck includes examination of: (B450.B.w28)

• Nasal cavity and larynx. (B450.B.w28)
• Oral cavity: mouth, tongue, palatal pits, teeth and pharynx. (B450.B.w28)
  • Check for lesions in the mouth indicative of Foot-and-Mouth Disease (Note: Details of the disease in elephants are given on the page Foot-and-Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA). (D286.2.w2)
  • Eggs from bot flies may be found around the teeth and the base of the tusks. See: Bots in Elephants
  • The elephant's age can be estimated from the molars. (P80.1.w1)
• Trunk: any swelling, trophy or injury. 
  • The trunk can be removed from the head at the trunk's base by cutting above and between the tusks. (P80.1.w1)
• Tusks: look for a black spot at the tip, broken tips, longitudinal cracks, irregularities of the dentine. (P80.1.w1)
• Cervical trachea and oesophagus. (B450.B.w28) (see also the sections on the respiratory tract and gastro-intestinal tract)
• Sensory organs: eyes, ears, Jacobson's organ, temporal glands and proboscis. (B450.B.w28, B411.IV.w4)
  • Dissect the eyes from the orbits. (D286.2.w2)
  • Heavy discharge from the temporal glands may indicate severe stress before death (e.g. from pain or disease), but is normal in bulls in musth (these bulls will also have urine staining on the inside of the hind legs, and a rank smell). (B411.IV.w4, P80.1.w1)
• Endocrine system: pituitary gland, thyroid and parathyroid glands. (B450.B.w28)
• Nervous system: brain, meninges and cervical spinal cord. (B450.B.w28)

Diseases affecting the trunk of elephants include

• Flaccid Trunk Paralysis in African Elephants Loxodonta africana

• Trunk Paralysis (Non-Specific) in Elephants

• Trunk Wounds in Elephants

Diseases affecting the mouth and tusks of elephants include

• Bacterial Tusk Infection in Elephants

• Foot-and-Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA

• Molar Maldevelopment and Malocclusion in Elephants

• Tusk Development Problems in Elephants

• Tusk Sulcus Infection in Elephants

• Tusk Injury in Elephants

Eye and ear diseases of elephants include

• Cataracts in Elephants

• Conjunctivitis in Elephants

• Ear Mites in Elephants

• Frostbite

• Iatrogenic Segmental Ear Gangrene in Elephants

• Keratitis and Corneal Ulceration in Elephants

• Miscellaneous Eye Lesions in Elephants

• Otitis Externa in Hedgehogs and Elephants

• Myiasis

Recommended tissue sampling (B450.B.w28)

• Gross lesions.
• Representative sections of the brain. (B450.B.w28)
• Pituitary gland, including dura. (B450.B.w28)
• Eyes. (B450.B.w28)
• Cross section of the tongue that include both mucosal surfaces. (B450.B.w28)
• Intact thyroid and parathyroid with a single transverse cut. (B450.B.w28)
• Section of thymus. (B450.B.w28)

Sampling for Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs and Elephants)

• Open the whole length of the trunk and check for any plaques, nodules or suspicious lesions. Any found should be divided into two sections, one into formalin and sent for histopathology, the other deep frozen and sent for culture. (See: D292 - Elephant Necropsy Protocol (Elephas maximus and Loxodonta africana))

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: DD293 - Elephant Research And Tissue Request Protocol

Eyes

• Slightly sunken orbits may indicate dehydration. (J1.35.w5)
• Note any ocular lesions: Ocular Disease in Bears
  • Ocular nematodes have been detected in bears. See: Eyeworms in Bears
  • An inactive hyperpigmented chorioretinal scar in the tapetal region of the left eye, compatible with a healed blastomycotic chorioretinal granuloma, was noted in a bear with blastomycosis (Blastomycosis in Bears)
  • Temporary unilateral corneal opacity has been seen following canine adenovirus 1 infection (Infectious Canine Hepatitis (with special reference to Bears))
  • Ocular neoplasms reported in bears include squamous cell carcinoma, melanoma of the eyelid and myxoma of the palpebral conjunctiva. (B16.9.w9, P1.2002.w5) See: Neoplasia in Bears
  • One case of a grass awn penetrating the cornea of a bear cub has been observed. (J417.20.w1, V.w93)

Ears

• Lesions associated with biting insects may be found on the tips of the ears. See: Biting-Blood-sucking and Nuisance Flies Affecting Elephants and Bears
• Grass awns and other plant material have been found commonly in the external auditory canals of Pacific Northwest Ursus americanus - American black bear. (J417.20.w1, V.w93)

Mouth, external nares, throat

• Check the condition of the tongue, teeth, gums and tooth sockets, also the salivary glands, larynx, trachea, thyroid glands, thymus and lymph nodes. (D247.7.w7)
• Check the mouth and the throat for foreign bodies and dental disease. See: 
  • Choke in Elephants and Bears
  • Dental Disease in Bears
  • Oral Foreign Bodies in Hedgehogs and Bears
• Check the mouth of cubs for signs of Candidiasis in Waterfowl and Bears (with notes on Elephants)
• Examination of the tongue musculature (at least in Ursus americanus - American black bear) may detect the presence of Gonglyonema pulchrum nematodes. See: Gonglyonema pulchrum Infection in Bears

Skull, internal nasal chambers

• Abnormalities of the skull and facial bones have been see with cretinism, hydrocephalus and severe rickets. See:
  • Calcium-Vitamin D Metabolism Imbalance (with special reference to Hedgehogs, Elephants and Bears)
  • Cretinism in a Bear
  • Hydrocephalus in Bears
• The brain, pituitary and meninges, and the nasal cavity, maxillary and frontal sinuses should be examined. (D247.7.w7)
  • Examination of these will be restricted if a cosmetic necropsy is being carried out so the skull can be preserved for taxonomic research. (D247.7.w7)
  • Examination of the nasal chambers and sinuses may reveal the presence of fluid and flukes. See: Prouterina wescotti Infection in Bears
  • Look for any abnormalities of the brain vascular system. See: Cerebral Arteriosclerosis in a Bear

• Note any abscesses. These may be associated with: (B600.17.w17)
  • Dental disease.
  • Foreign bodies.
  • Fight wounds.
  • Overgrown teeth penetrating the tissues.
• Wet fur under the chin may be present with excessive salivation due to cheek tooth spurs. (B600.17.w17)
• Epiphora and associated facial dermatitis may be associated with: (B600.17.w17)
  • Ocular disease.
  • Dental disease.
• Check particularly for dacrocystitis and conjunctivitis. (J72.49.w4)

Ears

• Note any wounds. (B611.4.w4, B614.5.w5)
• Note any foreign materials. (B611.4.w4, B614.5.w5)
• Check the ears thoroughly for signs of ear disease; (J72.49.w4) Inflammation of the ear canal, and presence of exudate/pus may be noted with: (B600.17.w17)
  • Bacterial Otitis Media - Interna in Lagomorphs
  • Psoroptes cuniculi - Ear Mite Infection in Lagomorphs
  • Aural foreign body.

Eyes

• Swelling of the eyelids is seen with:
  • Myxomatosis
  • Acute renal failure and associated fluid retention. (B600.17.w17)
• Purulent discharge:
  • Myxomatosis
  • Conjunctivitis in Lagomorphs.
  • Dacrocystitis in Rabbits.
  • Conjunctival foreign body. (B600.17.w17)
• Exophthalmos in Lagomorphs (B600.17.w17)
  • Trauma.
  • Retrobulbar abscess (Abscessation in Lagomorphs)
  • Retrobulbar tumour (Naturally Occurring Neoplasia in Lagomorphs)
  • Retrobulbar cyst (Tapeworm Cyst Infection in Lagomorphs)
  • Glaucoma in Rabbits

Nose

• Note any wounds. (B611.4.w4, B614.5.w5)
• Note any foreign materials. (B611.4.w4, B614.5.w5)
• Clear nasal discharge may be present with: (B600.17.w17)
  • Rhinitis in Rabbits
  • Pasteurellosis in Lagomorphs
• Purulent dischange may be present with: (B600.17.w17)
  • Nasal (and other URT) Foreign Bodies in Lagomorphs
  • Periapical infection (Facial Abscesses in Rabbits)
  • Myxomatosis (Viral Disease)
• Haemorrhagic discharge may be present with: (B600.17.w17)
  • Rabbit Haemorrhagic Disease
  • Trauma, including predator attack. (B600.17.w17)
    • Lacerations & Punctures, including bite wounds
• Purulent exudate in the nasal cavity may be associated with:
  • Nasal (and other URT) Foreign Bodies in Lagomorphs (hay seeds, hair plugs) (B600.17.w17)
  • Periapical abscess (maxillary cheek teeth). (Facial Abscesses in Rabbits)
  • Nasolacrimal duct dilatation and rupture (Dacrocystitis in Rabbits). (B600.17.w17)
  • Bacterial infection (Rhinitis in Rabbits)
  • Sinusitis.

Mouth

• Examine all surfaces including the buccal mucosa, tongue and teeth. (B611.4.w4, B614.5.w5)
• Note any wounds. (B611.4.w4, B614.5.w5)
  • Oral Soft Tissue Trauma in Rabbits
• Note any foreign materials. (B611.4.w4, B614.5.w5)
• Note any fractures of the jaw. (B600.17.w17)
• Note any dental abnormalities: (B600.17.w17) incise at the buccal commisures to allow thorough examination. (J72.49.w4) 
  • Congenital Incisor Abnormalities in Rabbits
  • Trauma-associated (fractured teeth) Acquired Incisor Abnormalities in Rabbits
  • Acquired. Acquired Molar Abnormalities in Rabbits
• Note any soft tissue ulceration associated with: (B600.17.w17)
  • Overgrown teeth
  • Penetrating foreign bodies
  • Iatrogenic (from dental burr or tooth clippers). 

To remove the tongue, larynx and associated structures: incise from the mandible down to the sternum. Reflect the skin to either side. Starting at the point of the chin, cut along the medial side of each mandible through the floor of the mouth. Using toothed forceps, grasp the tongue and pull caudally, while dissecting through the tissues of the throat and continuing posteriorly until the required tissues have been detached. (B611.4.w4, B614.5.w5)

Skull

• Palpate for any abnormalities. (B611.4.w4, B614.5.w5)
• If the skull does not need to be preserved, remove the parietal and nasal bones to allow examination of the nasal chambers, sinuses and brain. (B600.17.w17)
• The head can be split along the midline to examine the nasal cavities and sinuses. (J72.49.w4)
• Check the tympanic bullae for pus (Bacterial Otitis Media - Interna in Lagomorphs). (J72.49.w4)

• Check the mouth, nostrils, eyes and ears for any abnormalities or discharges. (B232.16.w16)

Conditions which may be seen in the mouth include:

• Dental Abscesses in Ferrets

• Fractured Canines in Ferrets

• Oral Foreign Bodies in Hedgehogs, Bears and Ferrets

• Periodontal Disease in Rabbits and Ferrets

• Persistent Deciduous Canines in Ferrets

• Salivary Mucocoele in Ferrets

• Supernumerary incisors (incidental).

• Note the colour of the mucous membanes, and any exudates.
• Examine the eyes and ears; note any exudates.
• Examine the oral cavity and pharynx.
• Examine the teeth.
• Examine the tongue.
• Examine the larynx and the laryngeal air sacs.
• Examine the salivary glands (mandibular, parotid).
• Examine the thyroids and parathyroids.
• Examine the cervical and cranial lymph nodes.
• Examine the oesophagus.

(D410)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Note the general appearance of carcass pelage (fur, spines, whiskers (vibrissae)) - e.g. condition, wet, muddy, oiled, clean, bloody, moult. 
  • If contaminated, note extent and areas affected.
  • If in moult, note the extent and take into account the season.
  • Missing patches of fur should be correlated with wounds, parasites, history of pruritus (itching), entanglement etc.
  • Condition of the fur may indicate whether the animal was in good condition or a state of debility prior to death.
• Note any wounds present and characterise them in terms of age, size, location, degree of sepsis.
• Check for the presence of external parasites (ectoparasites)- fleas, lice, ticks, maggots etc. 
  • Note the species, numbers (accurate or approximate indication), distribution.
  • Particular attention should be paid to checking the predilection sites for external parasites e.g. armpits (axillae), groin, perineum, hoof clefts, eyes, ears etc.
• Systematically part the hair over multiple areas of the body to examine the skin, looking for lesions including bite wounds, papules, macules, pustules, comedones, furuncles etc.
• Examine all body orifices, including the anus, vulva, prepuce, cloaca, marsupium (pouch) as appropriate, for evidence of haemorrhage, discharge, parasites, maggots (see Myiasis), abnormal growths etc.
• The carcass must be turned over allowing a full examination to be performed over both sides of the body.
• Examine the perineum and hindquarters for evidence of faecal scouring suggestive of diarrhoea (scour).
• At the end of the post mortem, the skin should be removed from all of the carcass to allow complete examination of the subcutaneous layers.
  • Check for evidence of bruises, wounds, parasites, ballistic wounds, snake bites, burns, etc.

Samples

• Collect external parasites in 70% ethanol for preservation and further identification as required.
• Skin samples for further examination include collection and microscopic (or hand lens) examination of scurf, scale, hair pluckings, deep skin scrapes, Sellotape (clear sticky tap) impression smears, Woods lamp (UV) examination for dermatophytes, swab or sample for microbiology (bacteriology, mycology), lesion sample for histopathology, lesion sample for virology.
• Tissue samples, swabs for culture and impression smears should be taken when tissues are first examined and before they become contaminated. 
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.

(B411.II.w2, B411.III.w3, P24.327.w13, D91.4.w1, V.w26)

• Closely examine the vulva, prepuce, perineum and any wounds for evidence of fly eggs or maggots, particularly in the summer. (See: Myiasis)
• Check all the spines and skin for evidence of wounds whose appearance may give an indication as to their cause:
  • Clean fresh lacerations, snout and limb amputations following strimmer injuries. (See: Lacerations & Punctures, including bite wounds)
  • Encircling snare-like wounds after being caught in plastic drink can trimmings or netting. (See: Foreign Body Entanglement & Snaring))
  • For information on prevention see: Garden Management for Hedgehogs (Erinaceus europaeus)
• Examine the spines for evidence of charring or burns, particularly in the autumn or if there is a history of garden fires in the area. (See: Burns)
• Examine between the spines for ectoparasites, particularly ticks (see: Tick Infection in Mammals) and fleas (see: Flea Infection in Mammals)).
  • Increased numbers of ectoparasites are often seen on hedgehogs which were debilitated prior to death.
• Check for evidence of scurf, scale and hair loss around the face, muzzle, ears, belly and 'skirt' (margin between the haired and spined areas).
  • Samples for dermatophyte culture and skin scrapings for parasitological investigation may be required for diagnosis.
  • See: Ringworm in Hedgehogs, Caparinia Mange of Hedgehogs
• Note any smells which may be associated with wounds, mite infection, other skin problems, or contamination. (B337.6.w6)

• In wild elephants, check carefully for penetrating wounds, for example due to bullets, tusks. (B411.IV.w4, D286.2.w2, P80.1.w1) or (particularly in calves) mauling by a large carnivore. (D286.2.w2)
  • In captive elephants, check for injuries which may have been inflicted by mahouts. (D286.2.w2)
• Look for signs of dislocations or fractures. (D286.2.w2)
• Skinning of the carcass may be required if any penetrating wounds are present (Wounds in Elephants), or to detect subcutaneous abscesses (Subcutaneous Abscess in Elephants). (P80.1.w1)
• Check for external parasites such as fleas or ticks. (P80.1.w1) (Flea Infection in Mammals, Tick Infection)

• Burns and Smoke Inhalation (with special reference to Waterfowl, Hedgehogs and notes on Elephants)

• Dermatitis in Elephants

• Dermoid Cysts in Elephants

• Elephant Foot Fly Infection

• Epidermoid Cysts in Elephants

• Filariasis of Elephants

• Flea Infection in Mammals (with special reference to Hedgehogs and a note on Elephants)
• Foot-and-Mouth Disease
  • (Note: Details of the disease in elephants are given on the page Foot and Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA)

• Foot Wounds in Elephants

• Frostbite in Waterfowl (with notes on Elephants)

• Iatrogenic Segmental Ear Gangrene in Elephants

• Idiopathic Ventral Oedema in Elephants

• Ingrowing Nails in Elephants

• Herpesvirus Infection in Elephants

• Hemorrhagic Filariasis Dermatitis of Elephants

• Hyperkeratosis in Elephants

• Lice Infection in Elephants

• Myiasis (with special reference to Waterfowl, Hedgehogs and notes on Elephants)

• Nail Abscess in Elephants

• Nail Cracks in Elephants

• Nail Overgrowth in Elephants

• Neoplasia in Elephants

• Papillomatosis (with special reference to Elephants)

• Pododermatitis in Elephants

• Poxvirus Infection in Elephants

• Ringworm in Hedgehogs (with notes on Elephants)

• Foot Wounds in Elephants

• Sole Abscess in Elephants

• Sole Cracks in Elephants

• Sole Overgrowth in Elephants

• Skin Myiasis in Elephants

• Skin Ulcers in Elephants

• Stephanofilarial Dermatitis in Elephants

• Subcutaneous Abscess in Elephants

• Tick Infection (with special reference to Hedgehogs)

• Trunk Wounds in Elephants

• Zinc Deficiency in Waterfowl (with special reference to Elephants)

• Wounds in Elephants

• Xeroderma in Elephants

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Full thickness of abdominal skin. (B450.B.w28)

Examination of the skin and fur of bears may reveal visible external parasites, including ticks, chiggers (visible as orange specks), fleas, lice and maggots. See:

• Chiggers (Trombiculiasis) (In Waterfowl, Hedgehogs and Bears)
• Flea Infection in Mammals (with special reference to Hedgehogs, Bears and a note on Elephants)
• Lice Infection in Elephants and Bears
• Myiasis (with special reference to Waterfowl, Hedgehogs, Elephants and Bears)
• Tick Infection (with special reference to Hedgehogs, Elephants and Bears)

Hair loss (alopecia) and/or crusting or scaling of the skin may be seen associated with various mange mite infections. Note: detection of mites other than chiggers requires examination of deep skin scrapings and/or skin biopsies. 

• Audycoptic Mange in Bears
• Biting-Blood-sucking and Nuisance Flies Affecting Elephants and Bears
• Demodicosis (with special reference to Hedgehogs and Bears)
• Sarcoptic Mange

Other skin conditions which may be apparent include:

• Bacterial Dermatitis in Bears
• Dermatophilosis in Bears
• Flea Infection in Mammals (with special reference to Hedgehogs, Bears and a note on Elephants)
• Greening of the Hair Coat in Polar Bears
• Hypothyroidism in Bears
• Lice Infection in Elephants and Bears
• Pityrosporum pachydermatis in a Bear
• Pythiosis in Bears
• Ringworm in Hedgehogs (with notes on Elephants and Bears)
• Skin neoplasia (Neoplasia in Bears)
• Unspecified skin conditions of Bears
• Vitamin A-responsive Skin Disease in Bears

• Skin discolouration, including postmortem lividity. (B611.4.w4, B614.5.w5)
• Any wounds. (B600.17.w17, B611.4.w4, B614.5.w5, J72.49.w4)
  • From fights with cage mates, attack by predator, or associated with recent dematting. (B600.17.w17)
    • Lacerations & Punctures, including bite wounds (with special reference to Waterfowl, UK Wildlife, Bears and Lagomorphs)
• Any ulcers, tumours or scars. (B611.4.w4)
• Fur condition. (B611.4.w4)
• Any abnormal fur.
  • Take samples for further examination. (J72.49.w4)
    • See: Clinical Pathology of Lagomorphs - Skin Testing
• Check for external parasites and for scale suggesting Cheyletiellosis (B600.17.w17, J72.49.w4) Ectoparasite infections which may be seen include:
  • Cheyletiellosis
  • Chiggers (Trombiculiasis) (In Waterfowl, Hedgehogs and Bears and Lagomorphs)
  • Demodicosis (with special reference to Hedgehogs, Bears and Lagomorphs)
  • Flea Infection in Mammals (with special reference to Hedgehogs, Bears and Lagomorphs, and a note on Elephants)
  • Lice Infection in Elephants, Bears and Lagomorphs
  • Miscellaneous Mites Reported on Wild Lagomorphs
  • Psoroptes cuniculi Ear Mite Infection in Lagomorphs
    • May be found around the anus and scrotum as well as in the ears. (J72.49.w4)
  • Red Fowl Mite Infection In Lagomorphs
  • Sarcoptic Mange
  • Tick Infection (with special reference to Hedgehogs, Elephants, Bears and Lagomorphs)
• Presence of blood. (B611.4.w4, B614.5.w5)
• Palpably enlarged superficial lymph nodes. (B611.4.w4, B614.5.w5)
• Examine the external genitalia for any abnormalities. (B611.4.w4, B614.5.w5)
• Note any abscesses (Abscessation in Lagomorphs). These may be associated with: (B600.17.w17)
  • Dental disease. (Facial Abscesses in Rabbits)
  • Foreign bodies.
  • Fight wounds (Lacerations & Punctures, including bite wounds (with special reference to Waterfowl, UK Wildlife, Bears and Lagomorphs)).
  • Note: abscesses on the limbs may be indicative of haematogenous spread of infection. (B600.17.w17)
• Perineal area wet/soiled, indicating: (B600.17.w17, B611.4.w4)
  • Urinary Incontinence in Rabbits
  • Caecotroph Accumulation in Rabbits
  • Poor husbandry.
• Diarrhoea leaking from the anus associated with:
  • Enterotoxaemia. (B600.17.w17) (Clostridial Enteritis and Enterotoxicosis in Rabbits)
  • Enteritis. (B600.17.w17)
    • Colibacillosis
    • Clostridial Enteritis and Enterotoxicosis in Rabbits
    • Rabbit Enteric Coronavirus Infection (unlikely)
    • Rotavirus Infection in Rabbits
    • Salmonellosis
    • Tyzzer's Disease
  • Intestinal Coccidiosis in Hedgehogs and Lagomorphs 
• Dermatitis around the anus associated with Treponematosis in Lagomorphs. (J72.49.w4)
• Mammary development due to pregnancy, lactation or pseudopregnancy (Pseudopregnancy in Rabbits). (B600.17.w17)
  • Consider whether there is mastitis (Bacterial Mastitis in Rabbits). (B600.17.w17)
• Take samples of abnormal skin for histopathological examination, particularly any lesions at mucocutaneous junctions. (J72.49.w4)

• Examine the skin and hair for any lesions. (B232.16.w16)
• If any abnormalities are detected, take hair and skin samples for further examination. (B232.16.w16) See Clinical Pathology of Ferrets - Skin Testing
• Bacterial infections include:
  • Abscessation in Lagomorphs and Ferrets
  • Bacterial Dermatitis in Lagomorphs and Ferrets
  • Bacterial Mastitis in Rabbits and Ferrets
• Fungal infections which can present with skin lesions include:

  • Blastomycosis (with special reference to Bears and Snow Leopards)

  • Coccidioidomycosis in Ferrets

  • Cryptococcosis in Ferrets

  • Malassezia Infection in Ferrets

  • Mucormycosis in Ferrets

  • Ringworm in Hedgehogs, Bears, Lagomorphs and Ferrets (with notes on Elephants and Snow leopards)

• External parasites which may be present include:

  • Chiggers (Trombiculiasis) (In Waterfowl, Hedgehogs and Bears, Lagomorphs and Ferrets)

  • Cuterebra (Warble) Infection in Lagomorphs (with notes on Ferrets)Demodicosis (with special reference to Hedgehogs, Bears, Lagomorphs and Ferrets)

  • Flea Infection in Mammals (with special reference to Hedgehogs, Bears, Lagomorphs and Ferrets, and a note on Elephants)

  • Foot rot (Sarcoptic Mange)

  • Hypoderma bovis (Warble) Infection in Ferrets

  • Lynxacarcus mustelae Fur Mite Infection in Ferrets

  • Myiasis (with special reference to Waterfowl, Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)

  • Otodectes cynotis Ear Mite Infection in Hedgehogs and Ferrets

  • Sarcoptic Mange (with special reference to Hedgehogs, Bears, Lagomorphs and Ferrets)

  • Seasonal Alopecia in Ferrets

  • Tick Infection (with special reference to Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)

• Look for signs of bites, other wounds and stings; wounds are not ucommon, partucularly on the necks of female ferrets following mating:
  • Arthropods Bites and Stings in Ferrets

  • Lacerations & Punctures, including bite wounds (with special reference to Waterfowl, UK Wildlife, Bears, Lagomorphs and Ferrets)

  • Snake Bite in Elephants and Ferrets

• Also look for lesions indicating:

  • Atopy in Rabbits and Ferrets

  • Canine Distemper with special reference to Ferrets and notes on Bears

  • Neoplasia in Ferrets

• Examine the hair and skin. Look for any evidence of trauma.
• Examine the external genitalia.
• In females, examine the mammary glands and nipples.
• In neonates, examine the umbilicus.
• Note any presence of diarrhoea.

(D410)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment
• Clinical Pathology of Lagomorphs - Skin Testing
• Clinical Pathology of Ferrets - Skin Testing

• Examine the limbs for evidence of fracture, dislocation, swelling, crepitus, deformity, lacerations, wounds (including snare wounds) etc.
• Use the other limb as a comparison for reference to help identify pathology.
• Palpate the muscle bulk over all limbs, using this as another indicator of body condition. 
  • Muscle wasting (atrophy) affecting a single limb, or muscles groups within a single limb, may indicate recent disuse, possibly due to a long-standing injury affecting the limb.
• Provided the carcass is not in rigor mortis, manipulate each limb and joint through its range of movement, comparing each side with one another, and systematically working from the bottom (distal) to the top (proximal) of the limb. 
  • Abnormal increased or restricted range of movement at a joint should direct further internal joint examination. (See: Musculo-skeletal system - Internal section on this page)
• Radiographic examination may be indicated if an abnormality is detected on palpation.

Samples

• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.

(B411.II.w2, B411.III.w3, P24.327.w13, D91.4.w1, V.w26)

See lower down the page for Musculo-skeletal System - Internal Examination

• Check all limbs for evidence of fractures, which are often compound in the hedgehog.
• Limb amputations, sometimes bilateral, may occur as a result of garden strimmer injury.
  • For information on prevention see: Garden Management for Hedgehogs (Erinaceus europaeus)
• Check all the digits for evidence of locally erosive abscesses and ulcers. 
  • These may occur as a result of fighting in group housed hedgehogs whilst in captivity, particularly affecting the outer digits.

• Check for any fractures or dislocations. (D286.2.w2)
  • Check carefully for limb fractures in elephants which have collapsed in an awkward position. (B411.IV.w4, P80.1.w1)
• See: 

  • Bone Fractures in Elephants

  • Capture Myopathy

  • Cuticle Overgrowth in Elephants

  • Degenerative Joint Disease in Elephants

  • Foot-and-Mouth Disease
    • (Note: Details of the disease in elephants are given on the page Foot and Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA)

  • Foot Wounds in Elephants

  • Heel Cracks in Elephants

  • Ingrowing Nails in Elephants

  • Joint Luxations in Elephants

  • Mycoplasma Infection (with special reference to Waterfowl, Hedgehogs and Elephants)

  • Nail Abscess in Elephants

  • Nail Cracks in Elephants

  • Nail Overgrowth in Elephants

  • Overworn Sole in Elephants

  • Pododermatitis in Elephants

  • Sole Abscess in Elephants

  • Sole Cracks in Elephants

  • Suprascapular paralysis in Elephants

  • Traumatic Musculoskeletal Swellings in Elephants

  • Trunk Paralysis in Asian Elephants Elephas maximus

Record the number of toenails on each foot. (D293)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Gross swelling of the limb and both subcutaneous and deep muscular crepitation were noted in a bear with clostridial myonecrosis (Clostridial Myonecrosis in Bears).
• Fractures may be visible or palpable: Fractures in Bears
• Note the condition of the claws. See: Claw Overgrowth in Bears

• Check the soles of the feet for Ulcerative Pododermatitis in Lagomorphs. (J72.49.w4)
  • This may be seen in animals which have suffered from debility, obesity, disability, immobilization or poor husbandry. (B600.17.w17)
• Staining/exudate on the medial surface of the forepaws may indicate:
  • Rhinitis in Rabbits. (B600.17.w17)
  • Pasteurellosis in Lagomorphs. (B600.17.w17)
  • Nasal (and other URT) Foreign Bodies in Lagomorphs. (B600.17.w17)
  • Possibly dental disease. (B600.17.w17)
    • Acquired Incisor Abnormalities in Rabbits
    • Acquired Molar Abnormalities in Rabbits
    • Congenital Incisor Abnormalities in Rabbits
• Limb bone fractures (Fractures in Rabbits) may be present following trauma, including predator attack. (B600.17.w17)
• Swelling of the hind leg distal to a ring may be caused by occlusion of the blood supply to the distal limb (due to too small a ring, or fur/debris trapped under the ring. (B600.17.w17)
  • Rings are placed above the hock of pedigree rabbits for identification. (B600.17.w17)

• Examine the feet. (B232.16.w16)
• Check the legs, note e.g. joint enlargement. (B232.16.w16)
• Check for:
  • Fractures in Ferrets
  • Neoplasia in Ferrets (particularly chordomas). (B602.9.w9)

• Check for any fractures or malformations.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Wet the fur thoroughly with soapy water or disinfectant in water to minimize debris which may contaminate the internal organs.
• Position the carcass laying on its back or side, dependent on size.
  • Small carcasses may be kept in place using metal pins secured into a wooden board.
• For examination of small to medium sized mammals, the carcass can be laid on its back:
  • Both hind limbs should be cut along their inner aspects over the groin deep into the hip joints, severing their articular connections, until both limbs can then be placed out to the side.
  • Both forelimbs should be cut through their inner aspects over the armpits (axillae), disarticulating the shoulders by cutting through the muscular attachments inside the shoulder blade (scapulae).
  • All four limbs, whilst still attached to the torso by soft tissue, should then fall to the side, providing a stable base for further examination.
• For examination of large mammals (e.g. antelope, large ruminants), the carcass should be laid on its side.
  • With equids (Equidae - Horses (Family)) (and other perissodactyls (Perissodactyla - Odd-toed ungulates (Order)), the carcass should be laid on its right side for best handling of the caecum and colon. (B273)
  • With ruminants (Artiodactyla - Even toed ungulates (Order)) the carcass should be laid on its left side so that the rumen does not obscure access to the abdomen.(B273)
  • The uppermost hind limb should be cut along its inner aspect over the groin deep into the hip joint, severing the articular connections, until the limb can then be placed out to the side.
  • The uppermost forelimb should be cut through its inner aspect over the armpit (axilla), disarticulating the shoulder by cutting through the muscular attachments medial to the shoulder blade (scapula).
• Examine the mammary glands, where present and note their state of development and activity.
  • Where mammary development is significant, the glands should be removed at this stage.
  • Visually examine cut surface of the gland and palpate for evidence of scarring, mastitis, etc.
• Slit the skin from the mandible to the pubic area in the ventral midline: nick the skin over the chest in the midline, continue the incision rostrally (forwards) to the chin and caudally (backwards) to the pubic bone, taking care not to damage underlying tissues.
• Continue the incisions by making cross incisions across the chest and abdomen as necessary.
• Reflect the skin from the neck, chest and abdomen, noting the amount of subcutaneous fat, colour of subcutaneous tissues, bruising, haemorrhage or other lesions
• Incise the abdominal wall along the midline from a central point, working caudally (backwards) towards the pubic bones and cranially (forwards) towards the base of the rib cage and diaphragm.
  • Take care to avoid opening the gastro-intestinal tract inadvertently when incising the abdominal wall. 
  • Collection of gas occurs post mortem within the gastro-intestinal system, stretching the intestinal loops and making accidental damage early in the examination increasingly likely.
  • Contamination of the abdominal cavity with gastro-intestinal contents early in the examination may obscure visualisation of gross pathology and interfere with bacteriological examination. 
• Incise the abdominal muscles transversely (across the body) just below the sternum and parallel to the base of the ribcage and continue to the level of the vertebral column. 
• Examine the integrity of the diaphragm, checking for hernia, rupture, etc.
• Use bone cutters, knife, saw, or similar dependent on carcass size, to cut the ribs on both sides through the costo-chondral junctions (weakest point at the junction between cartilage and bone). Lift the sternum  upwards and remove the rib cage by dissecting free remaining attachments at the thoracic inlet, cutting through the clavicles as necessary.
• The pubic bones should be sawed through to allow full examination of the pelvic organs.
• Examine the chest (pleural) and abdominal (peritoneal) cavities for evidence of haemorrhage, exudate, transudate, effusions etc. 
  • Note the colour, viscosity, turbidity and volume of any fluid present.
  • Retain samples of the fluid for further testing as required.
• Examine the lining of the chest (pleural) and abdominal (peritoneal) cavities for evidence of adhesions, parasitic cysts, tumours, inflammation, infection etc. 
• Examine the internal organs in situ before handling them; study their relative layout, size and colour.
• Comment on the amounts of abdominal and mesenteric fat deposits.
• Visually examine and palpate each organ for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger. 

Samples

• Tissue samples, swabs for culture and impression smears should be taken when tissues are first examined and before they become contaminated. 
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Sample diaphragm tissue for detection of Trichinella infection (Trichinellidae - (Family)) in carnivore Carnivora - Carnivores (Order) and wild pig species (Suidae - Pigs (Family)) as indicated, dependent on region.
• Samples for toxicology should be either wrapped in foil ( if the suspected toxin is organic), plastic wrap ( if the suspected toxin is metal) or glass (if suspected toxin is unknown).
  • Fat samples are important for toxicological investigation (e.g. organochlorines).
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)

(B411.II.w2, B411.III.w3, P24.327.w13, D91.4.w1, V.w26)

• Hedgehogs should be positioned on their back for post mortem examination.
  • Pinning the limbs out onto a wooden board is helpful for positioning.
  • Arranging a hedgehog into a spread-eagled position immediately after death (i.e. before rigor mortis sets in) may be useful if a post mortem examination is to be carried out. (B337.6.w6)

• Note: TB has not been detected in wild elephants. (B411.IV.w4)

Abdomen

• To open the abdominal cavity, make an incision vertically between the last rib and the tuber coxae, over the bulge of the abdomen, down to the ground, then extend this incision midventrally towards the sternum. Lift the skin and abdominal wall flap to examine the organs in situ before dissecting and removing them (meat hooks are useful tools). (B411.IV.w4, P80.1.w1)
• Or: Incise from the sternum to the pubis through the skin, subcutaneous tissue and muscle, then up from the pubis to the anterior border of the iliac crest, and forward along the lumbar vertebrae. (D286.2.w2)
• Examine the organs in situ before removing them. (B411.IV.w4)
• Note the location and orientation, relative size, shape, the nature of any contents of hollow organs, and any smells coming from the organs. (D286.2.w2)

Thorax

• The thoracic cavity dissection should be conducted ideally by two people. A ventral midline incision is made, one person holds the retractor and the other person dissects the skin. The sternum is exposed and the ribs are separated at the costochondral junction. Retractors are adjusted to hold the thoracic cavity open. (B450.B.w28, D292)
• Or: Dissect the skin and subcutaneous tissue along the vertebral column, then incise down the centre and use ropes to flap the skin out the way. Make deep incisions cranial to the first rib and caudal to the last rib; connect the incisions along the ventral midline with a deep incision. Cut the vertebral articulations of the ribs, and the costo-chondral junctions at the sternum. The ribs and muscles can then be pulled back using ropes to expose the thoracic cavity. (D286.2.w2)
• Or: Dissect the diaphragm from the thoracic wall and then dissect the lungs from the diaphragm and thoracic wall. (B411.IV.w4, P80.1.w1)
  • Removal of the thoracic wall is easier in small than in large elephants. (P80.1.w1)
• The heart, lungs and associated structures are best removed "en bloc". (B450.B.w28)
• Note the location and orientation, relative size, shape, the nature of any contents of hollow organs, and any smells coming from the organs. (D286.2.w2)

Record fat stores and abnormal fluids or contents in the thoracic and abdominal cavity. (B450.B.w28, D286.2.w2, J359.7.w2)

• The presence of one to three litres of straw coloured fluid in the abdominal cavity is normal. (B411.IV.w4, P80.1.w1)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D293 - Elephant Research And Tissue Request Protocol

• Starvation in Waterfowl and Seabirds (with notes on Hedgehogs and Bears)
• Obesity in Bears

Tacky viscera are indicative of dehydration. (J1.35.w5)

Jaundice may be noted in bears with:

• Hepatic and Bile Duct Neoplasia in Bears
• Infectious Canine Hepatitis (with special reference to Bears)
• Sarcocystosis in Waterfowl and Bears

Check the subcutaneous tissues; note the presence of:

• Abscess in Bears
• Sparganosis in Bears
• Nematodes: Dirofilaria ursi infection in Bears

Thorax

• Note the contents of the thoracic cavity, the position of the organs, the condition of the pleura, mediastinum and lymph nodes. (D247.7.w7)
• Chronic pleuritis was seen in a bear with Pasteurella Infection in Hedgehogs (with notes on Bears)
• Extensive granulomas were present on all pleural surfaces in a bear with Blastomycosis in Bears

Abdomen

• Note the contents of the abdominal cavity, the presence and extent of fat stores, the position of the diaphragm and the organs, the condition of the peritoneum, omentum and lymph nodes. (D247.7.w7)
• Ascites is common with intra-abdominal neoplastic lesions. (B16.9.w9, J423.69.w1)
  • Hepatic and Bile Duct Neoplasia in Bears
  • Neoplasia in Bears
• Chronic peritonitis was seen in a bear with Pasteurella infection (Pasteurella Infection in Hedgehogs with notes on Bears))
• Extensive granulomas were present on all peritoneal surfaces in a bear with Blastomycosis in Bears
• Nematodes may be present. See Dirofilaria ursi infection in Bears
• A worm found free in the abdominal cavity of a bear was identified as a kidney worm. See: Dioctophyme renalis Infection in Bears

Note:

• Examination of the diaphragm musculature may detect the presence of Gonglyonema pulchrum nematodes or Trichinella. See:
  • Gonglyonema pulchrum Infection in Bears
  • Trichinella Infection in Hedgehogs and Bears

• Or place the rabbit in right lateral recumbency, dislocating the left limbs to provide exposure to the body cavities. (J72.49.w4)
• The fat is usually white in rabbits; it is yellow in some individuals which have a genetic lack of a certain hepatic enzyme needed for xanthophyll metabolization. (B600.17.w17)
• The muscles of rabbits are quite pale. (B600.17.w17, J72.49.w4)
• Look for haemorrhages indicating trauma. (V.w5)

Starting at the second rib and moving posteriorly, cut through the costal cartilages near the costo-chondral junctions. Take care not to cut too deep, into the lungs. Use a blade to separate the sternum from the diaphragm. Lift the sternum.

• Examine the pleural cavities. 
• Take and smears or pleural fluid samples required. 

Cut through the cartilage of the first rib on either side and sever any remaining attachments so that the whole breast plate can be removed. 

• Examine the lungs and lymph nodes in situ.
• Use a gloved hand to feel round the lungs, noting and breaking any adhesions.

(B611.4.w4, B614.5.w5)

• Examine the abdominal cavity with the organs in situ, noting the volume and colour of any free abdominal fluid, the presence of any omental adhesions, and the position and size of the various organs (spleen, liver, kidneys, stomach, caecum, female genital organs, urinary bladder). (B611.4.w4, B614.5.w5, V.w5)
• Note the amount of fat present. (J72.49.w4)
• Note any abscesses (Pasteurellosis in Lagomorphs) or haemorrhages (Colibacillosis, Clostridum piliforme infection - Tyzzer's Disease in Lagomorphs). (J72.49.w4)

• Place the ferret in dorsal recumbency and use disinfectant over the carcass to reduce local pollution with fur/dander, and bacterial contamination. (B232.16.w16)
• If necessary, pin the feet to a dissecting board. (B232.16.w16)
• Make a ventral midline skin incision, mandible to pubis. (B232.16.w16)
• Reflect the skin away from the midline. (B232.16.w16)
• Take hold of the xiphisternum and cut through the costro-chondal junctions on either side to open the chest cavity, reflecting the sternum and ribcage anteriorly. (B232.16.w16)
• Examine the thoracic contents in situ. note the presence of any fluid in the thoracic cavity or pericardium. (B232.16.w16)
• Open the trachea; note any fluid present, and any inflammation. (B232.16.w16
• Using a midline incision, open the abdominal wall and reflect laterally. (B232.16.w16)
• Note any fluid in the peritoneal cavity. (B232.16.w16)
• Check the abdominal organs in situ. (B232.16.w16)
• To remove the thoracic viscera, cut across the oesophagus and trachea and lift these upwards, exteriorising the heart and lungs and breaking any connections to the thoracic cavity dorsally. Cut through the oesophagus as it exits the thoracic cavity, passing through the diaphragm. (B232.16.w16)

Thorax:

• Note any effusions, adhesions or haemorrhage.
• Note the amount and condition of mediastinal and coronary fat.
• Examine the thymus.
• Examine the lymph nodes.
• Note any obvious abnormalities of the heart, lungs, trachea or oesophagus.
• Fix a section of the thymus/anterior pericardium.
• Take one sternebra as a source of bone marrow. For bone marrow cytology, cut through the sternebra, make a touch imprint and allow to air dry. 

Abdomen:

• Note any effusions, adhesions or haemorrhage.
• Check the amount and condition of the omental, mesenteric and perirenal fat.
• Examine the abdominal aorta and caudal vena cava.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• First examine the heart and great vessels in situ looking at their relative layout (e.g. patent ductus arteriosus (PDA), persistent right aortic arch), size, shape and colour. 
• The organs of the chest are removed by cutting the trachea and great vessels as far cranial (forward) as possible. Applying gentle traction, carefully dissect back the 'pluck' containing the thymus (large if young, remnant may be identified in adult), heart and lungs.
  • Alternatively, dissect along the inside of the blades of the lower jaw, free the tongue, larynx, trachea and oesophagus and remove together with the 'pluck'.
• Examine the 'pluck' as a whole and then dissect each organ free for individual examination.
• Dissect out the heart, leaving a portion of the major blood vessels intact. 
• Visually examine and palpate each organ for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
• Examine the surface of the pericardium, incise the pericardial sac and remove from the surface of the heart.
  • Note the volume, colour, turbidity and viscosity of any fluid present. 
  • Note the presence of adhesions between the pericardium and the surface of the heart.
• Examine the overall shape and symmetry of the heart and its epicardial (outer) surface.
• Systematically dissect the heart using scissors to follow the path of blood through the heart; first examine the right side chambers, followed by the larger and thicker walled left side chambers.
  • Evaluate the heart muscle and relative thickness and size of the heart chambers; look for dilatation and hypertrophy.
  • Examine the endocardium (inner lining) of the heart and the valves for evidence of irregularities, plaques, thrombus formation, calcification, etc.
• Examination of the appearance of blood within the heart chambers can be used to help estimate the time since death.
  • Unclotted blood in the left ventricle is suggestive of recent death and examination pre rigor mortis.
  • Disintegrating blood clots within the left ventricle suggest that the animal has been dead for in excess of 24 hours and has passed through the period of rigor mortis.
• Cut through the heart in serial sections, noting the presence of any discolourations, parasitic cysts, growths, infarcts, scars, etc.
• Weigh and record organ dimensions (maximal length and width) as required.

Samples

• Tissue samples, swabs for culture and impression smears should be taken as tissues are examined and before they become contaminated. 
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Samples of blood from the heart chambers are commonly taken for bacteriological examination.
  • This sample should be taken before the heart is handled or incised to reduce contamination.
  • The surface of the heart should be seared with hot metal and a sterile swab or pipette used to collect the sample.
• Samples of clotted blood are taken from the left ventricle for serology.
  • Blood clots are centrifuged to collect serum samples for further testing.
• Tissue to be fixed for histopathology should be no more than 3-5mm thick, and thinner if congested. 
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases. (J1.32.w7)

(B411.II.w2; B411.III.w3, B273, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Atrio-ventricular valve problems such as endocarditis may be found. See: Congestive Heart Failure in Hedgehogs

• Cut the pericardium with sharp scissors, examine the surface and any contents. (D286.2.w2)
• Note the fat around the heart. This should be abundant, firm and light in colour in a healthy elephant. (P80.1.w1)
• Incise the heart along the coronary groove. Examine the valves, endocardium, papillary muscles and any other visible structures. (D286.2.w2)
• Record the heart's weight. (B450.B.w28)
• Check for any parasitic lesions of the heart muscle or vessels. (P80.1.w1)
• Check for arteriosclerosis of the aorta. (P80.1.w1)

• Arteriosclerosis in Elephants

• Cardiac Glycoside Toxicity in Elephants

• Congenital Cardiac Abnormalities in Elephants

• Schistosomiasis in Waterfowl (with notes on Elephants)

• Encephalomyocarditis (with special reference to Elephants)

• Filariasis of Elephants

• Trypanosomiasis in Elephants

• West Nile Virus Disease

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Longitudinal section including the atrium, ventricle and valves from both right and left heart and also the great vessels. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Note the position of the heart, the condition of the heart, pericardium and associated blood vessels. (D247.7.w7)
• Heartworms have been found in Ursus americanus - American black bears in North America; these have been incidental findings, without associated clinical signs. See: Heartworm Infection in Bears
• Valvular lesions have been found associated with a severe subacute fibrinosuppurative valvular endocarditis caused by a coagulase-positive Staphylococus aureus. See: Staphylococcosis (with special reference to Waterfowl, Hedgehogs and Bears)
• A wild adult female bear was found to have the congenital abnormality Fallot's tetralogy (Tetralogy of Fallot in a Bear)
• Check the aorta and other large arteries: 
  • Medial Calcification of the Aorta in Bears

• Note the amount and colour of pericardial fluid, and the presence of any haemorrhage or adhesions. 
• Pericarditis may be noted with Pasteurellosis in Lagomorphs. (J72.49.w4)

Lift the heart, cut the great vessels (vena cavae, aorta, pulmonary veins and pulmonary artery) and free the heart.

• Note: The heart and lungs can be removed together if preferred. (B611.4.w4, B614.5.w5)

The heart may appear abnormal with:

• Cardiomyopathy in Rabbits. (B600.17.w17)
• Congenital Cardiac Abnormalities in Elephants and Lagomorphs. (B600.17.w17)
• Tyzzer's Disease in Lagomorphs. (B600.17.w17)

Take a heart blood sample for serological testing if there is a suspicion of viral haemorrhagic disease (Rabbit Haemorrhagic Disease). (J72.49.w4)

Check for calcification of the large vessels (Soft Tissue Mineralization - Kidney Calcification in Rabbits). (J72.49.w4)

• Check for any external discolouration, adhesions, changes in texture or other visible/palpable lesions. (B232.16.w16)
• To examine the internal structures of the heart while preserving the endocardium and heart valves, incise from the left ventricular apex through into the left atrium and the aorta, and from the right ventricular apex up through the right atrium into the pulmonary artery. (B232.16.w16)

Consider:

• Dilated Cardiomyopathy in Ferrets
• Heartworm Infection in Bears and Ferrets (with a note on Lagomorphs)
• Hypertrophic Cardiomyopathy in Ferrets
• Valvular Heart Disease

• Examine the heart in situ, noting its position and any pericardial effusions or adhesions; if any effusion is present, collect for analysis and culture..
• Removing the heart and associated organs, examine the outside of the heart. Check the ligamentum arteriosus (ductus arteriosus) for patency.
• Check the position of the great vessels.
• Open the pulmonary arteries and check for thrombi.
• Separate the heart and aorta from the rest of the thoracic viscera. Note the presence of coronary fat, the external surface of the heart, particularly the coronary vessels, the relative filling of the atria and whether the heart was in distole or systole (contraction) at the time of death. Note the general shape of the heart; the apex should be relatively sharp.
• Measure the length of the heart (apex to top of atria)
• Measure the circumference around the coronary groove at the base of the atria.
• Open the right atrium, starting at the tip of the right auricle and opening the atrium parallel to the coronary groove and continuing into the vena cava. Remove the blood clot.
• Examine the atrio-ventricular valves and the foramen ovale.
• Following the caudal aspect of the septum, cut into the right ventricle; continue around the apex to the anterior side and then into the pulmonary artery. Removing post mortem blood clots, examine the endocardium (inner surface).
• Open the left atrium, starting at the auricle and continuing through to the pulmonary vein. Remove the blood clot and examine the valves.
• Following the caudal aspect of the septum, open the left ventricle, continuing round the apex to the anterior side and then into the aorta (remove blood as needed to give a clear view). 
• Open the thoracic aorta and examine this along its full length, intima and adventitia. Note the location and severity of any atherosclerotic or fibrous plaques. Section the aorta and preserve in formalin.
• Removing all post mortem clots, and washing if necessary in cool water or dilute formalin, examine the inner surface. Check the foramen ovale for patency.
• Separate the heart from the aorta, just behind the brachiocephalic arteries, and from the pulmonary artery and vena cava, close to the heart.
• Weight the heart (minus pericardium, great vessels and post mortem clots).
• Measure:
  • Height and circumference of the heart (see above)
  • Thickness of the right ventricular wall, left ventricular wall, interventricular septum.
  • Circumference of the right and levt atrioventricular valves, aortic valves, pulmonary valves.
• Spin down any "chicken fat" clots for serum, if required.
• Fix the entire heart or take sections of the following for histopathology:
  • Longitudinal sections of the left ventricle, right ventricle, atrioventricular valves, atria.
  • Myocardium, including coronary vessels, from the left evtricla and right ventrical.
  • Papillary muscle.
  • Septum, from the base of the atrioventricular valves (where the conduction system s found).
  • Ascending aorta just above the aortic valves (where dissecting aneurysms are most likely to be found).
  • Descending thoracic aorta.
  • Abdominal aorta.
  • Any lesions noted.
• Note: In addition to the whole length of the aorta, the iliac arteries and popliteal arteries should be examined as these are common sites of aneurysms in humans.
• Any fibrous or fatty streaks, any overt atherosclerosis, dissections, aneurysms and any other abnormalities should be noted (location, type, severity).

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• First examine the lungs, trachea (windpipe) and major bronchi (airways) in situ; study their relative layout, size, shape and colour.
• The organs of the chest are removed by cutting the trachea and great vessels as far cranial (forward) as possible. Applying gentle traction, carefully dissect back the 'pluck' containing the thymus (large if young, remnant may be identified in adult), heart and lungs.
  • Alternatively, dissect along the inside of the blades of the lower jaw, free the tongue, larynx, trachea and oesophagus and remove together with the pluck.
• Examine the pluck as a whole and then dissect each organ free for individual examination.
• Visually examine and palpate the lungs for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
  • Pooling of blood in the dependent lung under the effects of gravity (hypostatic congestion), should be recognised as an artifact.
• Using sharp scissors, cut along whole length of trachea (cut along both sides required, as tracheal rings are springy) into the major bronchi, noting presence of fluid, blood, fungal plaques, necrotic lesions, parasites, foreign bodies, etc.
• Cut through the lungs in serial sections, noting the presence of any water, froth, blood, fungal infection, abscesses, tumours, parasites, tuberculous lesions, etc.
  • Squeeze the lung tissue and observe for parasites within the airways.
• Placing a cut section of lung tissue within water may help evaluate congestion, consolidation, etc.
• Tracheo-bronchial lymph nodes should be examined visually, by palpation and on cut serial section for evidence of enlargement, focal lesions, abscesses, tumour metastases, tuberculous granulomata etc.
• Weigh and record organ dimensions (maximal length and width) as required.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases.(J1.32.w7)

(B411.II.w2, B411.III.w3, B273, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Close examination of the lungs for evidence of pneumonia associated with parasitic infection is essential.
  • Serial sections of the lung tissue should be made and then squeezed to see if their are any adult lung worms present within the major airways.
  • See: Lungworm Infection of Hedgehogs

• Cut along the primary, secondary and tertiary bronchi. Slice several times through any suspected lesion. (D286.2.w2)
• Submit any lung lesion for mycobacterial examination. (D286.2.w2)
• Submit the bronchial lymph nodes for TB culture even if they appear normal. (D286.2.w2)
• Look for any abnormalities affecting the nasal cavity, larynx, trachea, lungs and regional lymph nodes. (B450.B.w28)

Diseases of the respiratory system in elephants include:

• Aspiration Pneumonia in Elephants

• Cardiac Glycoside Toxicity in Elephants

• Coryzalike Syndrome in Elephants

• Choke in Elephants

• Interstitial Pulmonary Fibrosis in Elephants

• Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs and Elephants)

• Mycoplasma Infection (with special reference to Waterfowl, Hedgehogs and Elephants)

• Respiratory Nematode Infection in Elephants

Consider also the following which are not primary respiratory pathogens but may cause respiratory pathology:

• Anthrax (with special reference to Waterfowl and Elephants)

• Pasteurella Infection in Elephants

• Herpesvirus Infection in Elephants

Recommended tissue sampling (B450.B.w28)

• Gross lesions. 
• Sections from several lungs' lobes including a major bronchus. (B450.B.w28)
• Cervical, anterior, mediastinal and bronchial transverse sections. (B450.B.w28)

• Open the trachea and check for nodules or plaques. (D292)
• Check the tracheal and thoracic lymph nodes for caseous or granulomatous lesions. (D292)
• Palpate the whole of each lung for firm lesions. (D292)
• Any suspicious lesions found should be halved, one section being deep frozen and sent for culture, the other placed in formalin for histopathology. (D292)
• See: 
  • D292 - Elephant Necropsy Protocol (Elephas maximus and Loxodonta africana) [includes details of checking and sampling for TB in elephant necropsies]
  • D 303 - Guidelines for the control of tuberculosis in elephants 2003 [provides information on laboratories performing diagnostic tests for TB]

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D293 - Elephant Research And Tissue Request Protocol

• Note the position of the organs and the condition of the lungs, bronchi, respiratory blood vessels and associated lymph nodes. (D247.7.w7)
• Check the lungs, particularly the trachea, bronchi and bronchioles, for nematodes; both Crenosoma sp. and Capillaria aerophila lungworms may be found; these have been reported particularly in surveys of wild Ursus americanus - American black bears. 
  • Generally no pathology has been described associated with lungworms in bears, but viscous exudate in the air passages (trachea, bronchi, bronchioles) was described associated with Capillaria aerophila infection in one Ursus americanus - American black bear.
  • See: Lungworms in Bears
• Flukes have also been found in the lungs.
  • See: Prouterina wescotti Infection in Bears
• Lesions of TB have been found, very rarely, in the lungs of bears. (B214.3.4.w16)
  • See: Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs, Elephants and Bears)
• Lung lesions have been seen also with:
  • Aspiration Pneumonia in Elephants and Bears
  • Blastomycosis in Bears
  • Hepatozoon Infection in Bears
  • Hypersensitivity angiitis in a Bear
  • Respiratory Disease in Bears

• Examine the pleural cavities. 
• Take and smears or pleural fluid samples required. 

Cut through the cartilage of the first rib on either side and sever any remaining attachments so that the whole breast plate can be removed. 

• Examine the lungs and lymph nodes in situ.
• Use a gloved hand to feel round the lungs, noting and breaking any adhesions.

(B611.4.w4, B614.5.w5)

• Note: The heart and lungs can be removed together if preferred. (B611.4.w4, B614.5.w5)

Respiratory tract foreign bodies are common. (B600.17.w17)

Larynx:

• Foreign material such as hay. This may be inhaled or penetrated. (B600.17.w17)
  • See: Nasal (and other URT) Foreign Bodies in Lagomorphs
• Note: May be associated with dental disease. (B600.17.w17)
  • Acquired Incisor Abnormalities in Rabbits
  • Acquired Molar Abnormalities in Rabbits
  • Congenital Incisor Abnormalities in Rabbits

Trachea:

• Hyperaemia and oedema (J72.49.w4), or haemorrhagic exudate (B600.17.w17) is seen with
  • Viral haemorrhagic disease (Rabbit Haemorrhagic Disease). (B600.17.w17)

Lungs

• Haemorrhages (B600.17.w17)
  • Acute suppurative pneumonia, for example Pasteurellosis in Lagomorphs
  • Rabbit Haemorrhagic Disease
• Pneumonia (B600.17.w17)
  • Bacterial infections. See:
    • Pasteurellosis in Lagomorphs
    • Staphylococcosis (with special reference to Waterfowl, Hedgehogs, Bears and Lagomorphs)
  • Aspiration pneumonia
  • Mucoid Enteropathy in Rabbits
• Note: Lung pathology, often severe, may be an incidental finding at rabbit necropsy. (B600.3.w3)
• Take swabs of abscesses and discoloured areas of lung for bacterial culture. (J72.49.w4)
• Take impression smears for examination. (J72.49.w4)

• Open the length of the trachea, and bronchi; note any fluid present, and any inflammation. (B232.16.w16)
• Check for any external discolouration, adhesions, changes in texture or other visible/palpable lesions. (B232.16.w16)
• Take a sample of lung for bacterial culture. (B232.16.w16)

Consider:

• Aleutian Disease in Ferrets

• Aspiration Pneumonia in Birds, Elephants, Bears, Lagomorphs and Ferrets

• Bacterial Pneumonia in Rabbits and Ferrets

• Blastomycosis (with special reference to Bears and Ferrets)  

• Canine Distemper with special reference to Ferrets and notes on Bears

• Chlamydiosis - Psittacosis (with special reference to Waterfowl and notes on Bears and Lagomorphs)

• Coccidioidomycosis in Ferrets

• Heartworm Infection in Bears and Ferrets (with a note on Lagomorphs)

• Infectious Bovine Rhinotracheitis in Ferrets

• Influenza in Ferrets

• Neoplasia:

  • Lymphoma in Ferrets

  • Thymoma (see: Neoplasia in Ferrets)

• Pneumocystosis in Lagomorphs and Ferrets

• Pseudomonas Infection in Waterfowl, Lagomorphs and Ferrets (with notes on Hedgehogs)

• Examine the lungs, trachea and bronchi.
• Take and formalin-fix samples preferably from each lobe and certainly from one lobe on each side. If possible, inflate one lobe by infusing it under slight pressure with clean buffered formalin.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Examine the thyroid and parathyroid glands, the endocrine pancreas, and the adrenal glands, first in situ, then dissect free and examine in detail.
• Visually examine and palpate the organs for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
• Weigh and record organ dimensions (maximal length and width), as required.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases.(J1.32.w7)

(B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Transverse section across the adrenal gland, including cortex and medulla. (B450.B.w28)
• Entire pituitary gland including the dura. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Thyroid enlargement has been seen associated with Hypothyroidism in Bears and Cretinism in a Bear
• Thymomas have been seen:
  • Neoplasia in Bears
  • A thymoma was associated with Myasthenia Gravis in Bears

• Separate the pancreas from the small intestine when the GIT is removed from the abdomen. (B611.4.w4, B614.5.w5)
• The pancreas is diffuse and may be difficult to distinguish from mesenteric fat. (J72.49.w4)

• Carefully examine the adrenals for any enlargement or nodules indicative of Adrenocortical Neoplasia in Ferrets
• Check the pancreas for nodules perhaps 1-2 mm diameter which may indicate Insulinoma in Ferrets. (B627.12.w12, B631.30.w30) 
• Thymomas do occur in ferrets (Neoplasia in Ferrets). (J213.7.w5)
• Thyroid gland adenomas/adenocarcinomas are rare (Neoplasia in Ferrets). (B627.18.w18)

• Check the thyroid, thymus and pancreas. 
• Fix sections of thyroid, thymus and (with duodenum) pancreas. 
• Examine the adrenals.
• Weigh each adrenal.
• Take a longitudinal sample for histology from the left adrenal and a transverse section from the right adrenal; this allows identification of the two samples. N.B. use a very sharp blade to avoid crushing the tissue.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Ensure knowledge of the normal shape and number of liver lobes in the species under examination since significant variation exists between taxa.
• First examine the liver in situ; study its layout, size, shape and colour and relation to other abdominal organs.
• Dissect the liver lobes free of their attachments and remove.
• Visually examine and palpate the liver for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
• Note any haemorrhage on the liver surface or free in the body cavity; whether the capsule surface is intact or split; the presence of pale areas or other discolouration; whether lesions are flush with the surface, protruding or shrunken; whether edges of liver lobes are sharp (normal) or rounded (enlarged). 
• Weigh and record organ dimensions (maximal length and width), as required.
• Cut through the liver lobes in serial sections, noting the presence of any colour variations, size and shape of any lesions (pale areas, haemorrhages, abscesses, growths, cirrhosis, etc.).
• Note the size of the gall bladder; estimate the volume of bile present and describe its appearance. 
  • Note the degree of post mortem local bile staining of tissue.
  • Carefully express the gall bladder to determine whether any blockages may be present.
  • Dissect open the gall bladder and look for any material within the bile e.g. stones, parasites.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Samples for toxicology should be either wrapped in foil ( if the suspected toxin is organic), plastic wrap ( if the suspected toxin is metal) or glass (if suspected toxin is unknown).
  • Liver samples are important for toxicological investigation (e.g. organochlorines).
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)

(B273, B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Hepatic lesions may be seen occasionally associated with Leptospirosis. (J9.180.w1)
• Liver disease has been reported commonly as the main finding on post mortem examination of hedgehogs. (B291.12.w12)
  • See: Liver Disease in Hedgehogs

• Incise with a sharp knife. (D286.2.w2)
• Look for bile duct hookworms (Liver Nematode Infection in Elephants) and flukes (Liver Fluke Infection in Elephants). (B411.IV.w4, D286.2.w2, P80.1.w1)
• Check the bile ducts for calculi (gall stones). (P80.1.w1)
  • Cholelithiasis in Elephants
• Note: there is no gall bladder in elephants. The main duct is large. (B10.49.w21, B384.3.w3, P80.1.w1)
• Look for any other abnormalities (B450.B.w28), including hydatid cysts (B453.7.w7) (Hydatid cysts in Elephants)
• Record the weight of the liver. (B450.B.w28)
  • The liver may weigh as much as 68 kg in a large male. (P80.1.w1)

Hepatic diseases in elephants include:

• Cholelithiasis in Elephants

• Hydatid cysts in Elephants (rare)

• Liver Fluke Infection in Elephants

• Liver Nematode Infection in Elephants

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Sections from three lobes of the liver including the capsule. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Liver pathology (swelling, mottled appearance) may be seen in bears with hepatic sarcocystosis. See: Sarcocystosis in Waterfowl and Bears
• Liver lesions including necrosis have been seen with toxoplasmosis (Toxoplasmosis in Waterfowl (with notes on Hedgehogs, Elephants and Bears))
• Hepatic and Bile Duct Neoplasia in Bears is common in older captive bears.
• In bears from bear bile farms, Gall Bladder Fistula in Bears and associated Gall Bladder Lesions in Bears may be found.

• Once the abdomen is opened, note the position and the size of the liver. (B611.4.w4, B614.5.w5)

To remove the liver, sever the associated vessels and lift the liver. (B611.4.w4, B614.5.w5)

• Hepatic Lipidosis in Rabbits is common in the end-stage of many diseases; it develops rapidly if the rabbit does not eat (particularly in obese rabbits). (B600.17.w17)
• Hepatomegaly:
  • Rabbit Haemorrhagic Disease - pale yellow discolouration of the liver may be noted. (J72.49.w4)
  • Septicaemia
  • Hepatic Coccidiosis in Rabbits (also bile ducks thickened and enlarged). (B600.17.w17)
    • Impression smears of liver: may show various stages of the parasite. (B600.10.w10)
    • Wet smears of bile may show oocyts. (B600.10.w10)
  • Neoplasia.
  • Congestive Heart Failure in Rabbits
  • Lymphoma (enlared and pale) (Multicentric Lymphoma in Rabbits).
  • Toxoplasmosis in Waterfowl and Lagomorphs (with notes on Hedgehogs, Elephants and Bears)
• Nodules or foci present with bacterial infections and coccidiosis.
  • Hepatic Coccidiosis in Rabbits - impression smears from the cut surface can be examined for the presence of coccidia. (J72.49.w4)
  • Salmonellosis
  • Tyzzer's Disease in Lagomorphs
    • Causes miliary foci. Also GIT pathology - see below. (B600.17.w17)
  • Yersiniosis
  • Tularemia
  • Neoplasia (Naturally Occurring Neoplasia in Lagomorphs)
• Take impression smears for examination. (J72.49.w4)
• Histopathological examination is required for the diagnosis of hepatopathies such as Aflatoxicosis. (J72.49.w4)
• Liver samples can be submitted for analysis of lead concentration if Lead Poisoning is suspected.

• Cut through the liver lobes, check for any changes in architecture. (B232.16.w16)
• Take a sample of liver for bacterial culture. (B232.16.w16)
• Hepatomegaly (and splenomegaly and mesenteric lymphadenopathy) may be present with Aleutian Disease in Ferrets. B627.15.w15, J213.8.w3)
• Lesions may be found with:

  • Lymphoma in Ferrets (B602.3.1.w3a)

  • Other Neoplasia in Ferrets. (B602.3.1.w3a)

  • Toxoplasmosis in Waterfowl, Lagomorphs and Ferrets (with notes on Hedgehogs, Elephants and Bears)

• Weigh the liver.
• Note any liver lesions.
• Take sections with at least one section including bile ducts and gall bladder, as well as a section from at least one other lobe.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Ensure knowledge of the normal shape of the spleen in the species under examination since significant variation exists between taxa.
• Examination of the spleen is recommended early in the post mortem after the abdomen is cut open.
  • If the spleen is enlarged, with a black and tarry bloody appearance, testing for anthrax infection should be performed before progressing further. (See: Anthrax)
• First examine the spleen in situ; study its layout, size, shape and colour and relation to other abdominal organs.
• Dissect the spleen free of its attachments and remove.
• Visually examine and palpate the spleen for evidence of pathology; consider colour, consistency and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
• Note any haemorrhage on the surface of the spleen or free in the body cavity; whether the capsule surface is intact or split; the presence of pale areas or other discolouration; whether lesions are flush with the surface, protruding or shrunken; whether edges of the spleen are sharp (normal) or rounded (enlarged). 
• Weigh and record organ dimensions (maximal length and width), as required.
  • Splenic dimensions can be artifactually increased if the animal was euthanased with barbiturates.
• Cut through the spleen in serial sections, noting the presence of any colour variations, size and shape of any lesions (pale areas, haemorrhages, abscesses, growths, etc.).

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases. (J1.32.w7)

(B273, B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Splenic lesions may be seen occasionally associated with Leptospirosis. (J9.180.w1)

• The normal spleen is long and strap-like, dark blueish-red and covered by a tough, whitish connective tissue capsule. Dark red raised nodules looking similar to subcapsular haemorrhages are normal. A smaller extra "daughter spleen" is found occasionally. (B411.IV.w4, P80.1.w1)
• Check for congestion, which may indicate acute bacterial or protozoal disease. (D286.2.w2)
  • Anthrax
  • Trypanosomiasis in Elephants

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Cross section of the spleen including the capsule. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• The spleen may be affected by abdominal neoplasias, including hepatic neoplasia. See: Neoplasia in Bears, Hepatic and Bile Duct Neoplasia in Bears.
• Splenic laceration has been see as a darting injury. (P9.2004.w4)
• Gnathosoma sp. larvae have been found in the spleen. See: Gastro-Intestinal Nematode Infection in Bears.
• With Anthrax, in many species typically the spleen is grossly enlarged, dark and soft (pulpy). In a bear with anthrax, the spleen was not enlarged, blue-grey in colour, medium consistency, with a small quantity of dark red thickened blood. (J142.19.w1) Note: If anthrax is suspected the carcass should NOT be opened. Take peripheral blood smears and do not continue with the necropsy if a diagnosis of anthrax is made.
• Severe congestion of the spleen may be seen associated with clostridial disease. See: Clostridial Myonecrosis in Bears.
• The spleen may be displaced, enlarged and congested in bears with Acute Gastric Dilatation in Bears.
• Mild enlargement of the spleen was seen in a bear with Leptospirosis. (B16.9.w9)

• Note the position and size of the spleen. (B611.4.w4, B614.5.w5)

The spleen can be elevated from the stomach and the hilar vessels severed to separate it from the stomach. (B611.4.w4, B614.5.w5)

• The spleen is small, only up to 4 cm long and less than 1 cm diameter. (B600.17.w17, J72.49.w4)
• Splenomegaly may be present with: (B600.17.w17)
  • Rabbit Haemorrhagic Disease
  • Septicaemia e.g. Salmonellosis
  • Congestive Heart Failure in Rabbits
  • Toxoplasmosis
• Miliary punctate abscesses on the spleen (and the liver) suggest Yersiniosis. (J72.49.w4)
• Take impression smears for examination. (J72.49.w4)

• Note the size of the spleen and any obvious lesions.

  • Splenomegaly is a common finding in ferrets over about three years of age and can be considered a normal variation if there are no other findings indicating a problem.

• Lesions are seen with:

  • Lymphoma in Ferrets

  • Haemangioma / haemangiosarcoma (See: Neoplasia in Ferrets)

  • Hepatomegaly, splenomegaly and mesenteric lymphadenopathy may be present with Aleutian Disease in Ferrets. B627.15.w15, J213.8.w3)
  • Mammalian Tuberculosis ( lesions also may be found in the liver, lymph nodes, peritoneum and brain). (J195.11.w2)

• Examine the spleen.
• Weigh the spleen.
• Take thin sections for fixation,particularly if it is congested, as penetration of the formalin will be reduced. 

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Ensure knowledge of the normal shape of the kidneys in the species under examination, since significant variation exists between taxa.
• Ensure knowledge of the normal shape of the uterine tract in the species under examination, since significant variation exists between taxa.
  • e.g. uterus simplex, uterus bicornuate, uterus duplex.
• First examine the urinary system and reproductive system in situ; study its layout, size, shape and colour and relation to other abdominal organs.
• Trace the flow of urine through the urinary system in situ; follow the ureters from the kidney to the bladder and the urethra as possible.
  • Check for developmental abnormalities e.g. ectopic ureter. 
• Dissect the kidneys free of their attachments and remove.
• Note the amount of fat deposits present over the kidneys.
• Visually examine and palpate the kidneys for evidence of pathology; consider colour, consistency (firm, soft, pulpy) and shape in every case.
  • Organs should be palpated methodically by carefully feeling the tissue between the thumb and forefinger.
• Note whether the capsule is adherent or easily detaches from the kidney surface.
• Examine the cut surface of serial sections of both kidneys in two planes (longitudinal and transverse).
  • Study the distinction between the cortex and medulla. 
  • Note the presence of any cysts, stones (uroliths), parasites, etc.
• Weigh and record organ dimensions (maximal length and width), as required.
• Dissect open the bladder, examine its mucosa and contents for abnormalities.
• Examine the external genitalia including the vulva, prepuce, penis and testes (may be best performed earlier in the post mortem examination).
• Female reproductive tract:
  • Ovaries: size, whether ovaries are active (follicles present and growing, corpora lutea) or inactive, appearance of oviducts. Examine serial cut sections of both ovaries.
  • Dissect the tract open from the vulva through to the tips of the uterine horns. 
  • Note the presence and number of fetuses (estimate the stage of gestation, mummification), implantation scars; presence of retained placentae; evidence of inflammation, infection or growths.
• Male reproductive tract:
  • Examine the contents of the inguinal canal; check for hernia.
  • Testes: size, shape, colour, growths, active or regressed. Examine serial cut sections of both testes.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Samples for toxicology should be either wrapped in foil (if the suspected toxin is organic), plastic wrap ( if the suspected toxin is metal) or glass (if suspected toxin is unknown).
  • Kidney samples are important for toxicological investigation.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)

• Bladder stones and urinary tract infections have been recorded in hedgehogs. (B337.3.w3) See: 
  • Urinary Tract Infection in Hedgehogs
  • Urolithiasis in Hedgehogs
• Renal failure may occur in elderly Erinaceus europaeus - West European Hedgehog and in Atelerix albiventris - Four-toed hedgehog as young as two years old. See: Kidney Failure in Hedgehogs
• A full bladder may be present in individuals with spinal injuries or in young orphaned hoglets which have not been toileted by their mother or while in care.(B337.3.w3)
  • See: Fractures in Hedgehogs, Hand-rearing Hedgehogs (Erinaceus europaeus)
• Urinary retention has been noted in end-stage Hedgehog Paralysis associated with Demyelination. (J3.143.w5)
• Urine may contain Leptospira organisms (See: Leptospirosis (with a note on Hedgehogs)). (J9.180.w1)

Kidneys

• Cut through and strip off the capsule. (D286.2.w2)
• Cut the kidney in half lengthways and examine the cut surfaces. (D286.2.w2)
  • In the healthy kidney the junction between the cortex and medulla is clearly visible. (P80.1.w1)
• Cut through any suspected lesion. (D286.2.w2) See:
  • Renal Disease in Elephants
  • Pyelonephritis in Elephants

Bladder & urethra

• The urine should be a light straw-colour, is often turbid, but does not have a pronounced odour. (P80.1.w1)
• The bladder may be emptied during the elephant's death-throes; an empty bladder may indicate a slow death. (P80.1.w1)
• Check the bladder and urethra for lesions.
  • Cystitis in Elephants
  • Urethritis in Elephants

Look for any abnormalities affecting the reproductive system: in males, the testes, penis, prepuce and accessory glands; in females the ovaries, uterus, vagina, clitoris, mammary glands and (in pregnant females) placenta. (B450.B.w28)

• See:
  • Abortion in Elephants
  • Acyclicity in Elephants
  • Carbon Monoxide Toxicity in Elephants
  • Cervical Cysts in Elephants
  • Cervical Polyps in Elephants
  • Cystitis in Elephants
  • Dystocia in Elephants
  • Endometrial Cystic Hyperplasia and Polyps in Elephants
  • Lymphoid Follicular Vulvitis in Elephants
  • Metritis in Elephants
  • Neoplasia in Elephants
  • Ovarian Cysts in Elephants
  • Placentitis in Elephants
  • Priapism in Elephants
  • Poxvirus Infection in Elephants
  • Perineal Hernias in Elephants
  • Retained Placenta in Elephants
  • Stillbirth in Elephants
  • Testicular Abnormalities in Elephants
  • Urethritis in Elephants
  • Urinary Calculus in Elephants
  • Uterine Leiomyomas in Elephants
  • Vaginal Cysts in Elephants
  • Vaginal Prolapse in Elephants
  • Vestibular-Vulval Lesions in Elephants

Recommended tissue sampling (B450.B.w28)

• Gross lesions. 
• Sections from both kidneys that include cortex, medulla and pelvis. (B450.B.w28)
• Cross section of the bladder and a two cm section of the tubular structures. (B450.B.w28)
• Transverse section of the uterus. (B450.B.w28)
• One ovary or testis. (B450.B.w28)
• Representative samples from accessory sex organs. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Renal and retroperitoneal haemorrhage has been seen associated with a severe subacute fibrinosuppurative valvular endocarditis caused by a coagulase-positive Staphylococus aureus. See: Staphylococcosis (with special reference to Waterfowl, Hedgehogs and Bears)
• A renal tuberculous lesion was seen in one Ursus maritimus - Polar bear. See: Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs, Elephants and Bears)
• Congenital absence of one kidney has been noted: Kidney Agenesis in Bears
• Nephritis in Bears
• Oiling may cause renal damage.
• Neoplasia such as kidney adenoma has been seen. See: Neoplasia in Bears

To remove the urogenital system, cut behind each kidney and adrenal, freeing them from retroperitoneal tissues. Open the aorta and probe each renal artery, then cut through the renal vessels. Lift each kidney in turn and note the course of the ureters. Lift out and remove the kidneys and urinary bladder. (B611.4.w4, B614.5.w5)

Kidneys

• Enlarged kidney(s):
  • Septicaemia
  • Congestive Heart Failure in Rabbits
  • Neoplasia (e.g. pale and irregular with lymphoma). (B600.17.w17)
    • Multicentric Lymphoma in Rabbits
    • Naturally Occurring Neoplasia in Lagomorphs
• Pitting may be visible from Encephalitozoonosis in Lagomorphs; (B600.17.w17, J72.49.w4) this can be an incidental finding (i.e. no clinical signs were seen). (B600.17.w17)
• Serosal haemorrhage:
  • Rabbit Haemorrhagic Disease
  • Septicaemia.
  • Anticoagulant Rodenticide Toxicity (B600.17.w17)
  • Triazines. (B600.17.w17)
• Infarcts with Staphylococcosis. (B600.17.w17)
• Urolithiasis in Lagomorphs: (B600.17.w17)
• Renal parenchyma calcification: (B600.17.w17)
  • Chronic Renal Failure in Lagomorphs
  • Soft Tissue Mineralization - Kidney Calcification in Rabbits Vitamin D toxicity.
• Fatty infiltration:
  • Terminal event associated with Hepatic Lipidosis in Rabbits in many diseases causing prolonged anorexia. (B600.17.w17)
• Subcapsular cysts
  • Incidental finding of Renal Cysts in Rabbits. (B602.18.w18)
• Take impression smears for examination. (J72.49.w4)

Bladder

• Normal urine may vary in colour from pale yellow to orange, red or brown. (B600.17.w17)
• For further details on normal and abnormal characteristics of urine see: Clinical Pathology of Lagomorphs - Urinalysis
• The presence of calcium carbonate sediment is common, but excessive sludging is abnormal. (B600.17.w17)
• Uroliths - Urolithiasis in Lagomorphs

Genital tract

• Endometrial Hyperplasia - Uterine Polyps in Rabbits
• Uterine masses:
  • Endometrial Adenocarcinoma and other Uterine Neoplasia in Lagomorphs (Miscellaneous Disease Summary)
• Broad ligament containing necrotic masses:
  • Metastases from uterine adenocarcinoma..
• Endometrium contains blood-filled protrusions:
  • Endometrial Venous Aneurysms in Rabbits. 
• Dead fetuses in uterus:
  • Pregnancy Toxaemia in Lagomorphs
  • Metritis (Uterine Infection in Lagomorphs) associated with infections such as Listeriosis or Pasteurellosis in Lagomorphs.
• Pyometritis or abscesses Abscessation in Lagomorphs, Uterine Infection in Lagomorphs), Pasteurellosis in Lagomorphs
• Orchitis and Epididymitis in Rabbits:
  • Fight wounds - see Lacerations & Punctures, including bite wounds
  • Myxomatosis

(B600.17.w17, J72.49.w4)

• Incise the kidneys longitudinally, examining the internal structure. (B232.16.w16)
• Peel off the external capsule. (B232.16.w16)
• Examine the whole of the urogenital tract. (B232.16.w16)

Look for lesions of:

• Acute Renal Failure in Lagomorphs and Ferrets

• Bacterial Cystitis in Ferrets

• Bacterial Mastitis in Rabbits and Ferrets

• Ibuprofen Toxicity in Ferrets

• Hydronephrosis in Ferrets

• Hyperoestrogenism in Ferrets

• Neoplasia in Ferrets

• Polycystic Kidney Syndrome in Rabbits and Ferrets

• Prostatic Cysts in Ferrets

• Pyelonephritis in Elephants, Lagomorphs and Ferrets

• Pyometra in Ferrets

• Urolithiasis in Lagomorphs and Ferrets

• Vaginitis in Ferrets

• Zinc Toxicity in Waterfowl and Ferrets (Kidneys large, pale and soft; sometimes small capsular surface depressions). (J83.15.w4)

Urinary tract

• Examine the kidneys, ureters, bladder and urethra.
• Weigh each kidney.
• Take a longitudinal sample for histology from the left kidney and a transverse section from the right kidney; this allows identification of the two samples.

Reproductive tract

• Examine the reproductive tract., including the ovaries and uterus in females, testes, penis and accessory sex glands in males.
• In females, weigh each ovary and note three dimensions of the uterus.
  • If the female is pregnant, weight the placenta, note the side of the placental disc and the length of the umbilical cord. Take sections form the periphery and centre of the placental disc and from extraplacental fetal membranes. Weigh and measure the fetus and perform a post mortem examination on the fetus. Take sections of major organs from the fetus.
  • Take transverse sections of each part of the tract, including the ovaries.
• In males, weigh each testis and note the length and diameter. Note the length and diameter of the penis.
• Section the prostate (with urethra), seminal vesicles and testes transversely for fixing.

(D408)

• Clinical Pathology of Lagomorphs - Urinalysis
• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Examine the gastro-intestinal tract in situ; note any distension, abnormal layout (twisting or volvulus), discolouration, haemorrhage or lesions on the serosal surface.
  • Barbiturate crystals may be seen on the surface of the intestines if the mammal has been euthanased by intraperitoneal injection.
• Ensure knowledge of the anatomy of the gastro-intestinal system and the primary site of microbial fermentation; distinguish between simple stomached, foregut and hindgut fermenters.
• Tie off the oesophagus above or at the level of the diaphragm and the rectum as low as possible; cut above and below these ties respectively, carefully dissect the whole gastro-intestinal tract free of its mesenteric attachments.
• Carefully dissect through the soft tissues of the intestinal coils to release them and lay the tract out as one continuous strip.
• Samples for bacteriological examination should be taken before further handling of the tract to limit contamination; the surface of the area to be sampled should be seared with hot metal and a sterile swab or pipette introduced to take the sample material.
• Clips or ties can be used to isolate the contents of different sections of the gastro-intestinal tract; for example at the pylorus to isolate the gastric contents, at the ileo-caecal valve to isolate the small intestinal from the large intestinal contents.
  • Collection and examination of the gut contents from each section of the gastro-intestinal tract is particularly useful for parasitological examination.
  • Quantitative estimates of numbers of parasites present can be made with further techniques.
• Systematically examine the entire gastro-intestinal tract, usually from oesophagus to rectum, including the pancreas. Use scissors or an enterotome to open up the tract.
  • Check for ulcers, inflammation, haemorrhage, tumours.
• Comment on the consistency, colour and volume of the gut contents.
  • Note whether the mammal has eaten recently and what, if possible, it was feeding on.
  • Note any foreign bodies or parasites present
  • Note whether formed faecal pellets are present in the rectum.
• Examine the mesenteric lymph nodes visually, by palpation and on cut serial section for evidence of enlargement, focal lesions, abscesses, tumour metastases, granulomata etc.
• In the equidae (Equidae - Horses (Family)), particular attention should be paid to examination of the blood vessels (abdominal aorta, anterior mesenteric artery) to the large colon and caecum for the presence of thrombi, aneurysms or scars caused by parasites.

Samples

• Tissue samples, swabs for culture and impression smears should be taken when tissues are first examined and before they become contaminated. 
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)
• Fresh samples of contents and mucosal scrapings of duodenum, ileum, caeca may be examined microscopically for parasites, parasitic ova and coccidial oocysts.
• Samples of gastric contents should be taken when investigating a potential poisoning case.

(B273, B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• There are a number of gastro-intestinal parasites in hedgehogs. See: 
  • (Thorny-headed Worm) Acanthocephala Infection of Hedgehogs
  • Intestinal Coccidiosis in Hedgehogs
  • Cryptosporidiosis in Mammals (with special reference to Hedgehogs)
  • Giardia Infection in Mammals (with special reference to Hedgehogs)
  • Hedgehog Intestinal Nematode (Capillaria) Infection
  • Hedgehog Intestinal Fluke Infection
  • Hedgehog Nematode (Oesophagus & Stomach) Infection
• Bacterial infections which may be seen in hedgehogs include Salmonellosis and in hoglets Colibacillosis.

• Examine a faecal bolus from the rectum: the presence of large pieces of undigested food items may indicate problems chewing due to molar abnormalities (Molar Maldevelopment and Malocclusion in Elephants), old age, or an abnormal diet. (B411.IV.w4)
• Stomach:
  • Open the stomach (normally about 100-140 cm long and 40 cm diameter) and check the contents to see what has been eaten (e.g. mud in wild elephants in dry periods, for its moisture), the appearance of the contents (e.g. large pieces indicating tooth problems) (Molar Maldevelopment and Malocclusion in Elephants) and whether parasites are present (Parabronema spp. - see Elephant Gastro-Intestinal Nematode Infection). (B411.IV.w4, P80.1.w1)
  • Wash the contents out with water and examine the mucosa; look for parasites and their lesions. Parasitic granulomas and focal ulceration may be seen with Parabronema infection. (D286.2.w2) See:
    • Bots in Elephants
    • Elephant Gastro-Intestinal Nematode Infection
• Intestines:
  • These may have a total length of 18 m. (P80.1.w1)
  • Cut open longitudinally and examine the mucosa. (D286.2.w2)
  • The intestinal contents must be emptied out using a shovel and water to allow the mucosa to be examined. (P80.1.w1)
  • Section through any lesions. (D286.2.w2)
  • Examine for the presence of parasites. (D286.2.w2) Large numbers of parasites are commonly present in wild elephants. (P80.1.w1) See: 
    • Cestodes in Elephants
    • Elephant Gastro-Intestinal Nematode Infection
    • Intestinal Fluke Infection in Elephants

Gastro-intestinal diseases in elephants include

• Bots in Elephants

• Choke in Elephants

• Clostridium perfringens Infection in Elephants

• Colibacillosis (with special reference to Waterfowl, Hedgehogs and Elephants)

• Colic in Elephants

• Constipation in Elephants

• Elephant Gastro-Intestinal Nematode Infection

• Foreign Body Ingestion in Waterfowl (with notes on Elephants)

• Intestinal Fluke Infection in Elephants

• Salmonellosis (with special reference to Waterfowl, Hedgehogs and Elephants)

• Schistosomiasis in Waterfowl and Elephants)

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Sections three centimetres long of the oesophagus, stomach (cardia, antrum, pylorus), duodenum, jejunum, ileum, caecum, colon and omentum. (B450.B.w28)
• Transverse cut of the mesenteric lymph node. (B450.B.w28)
• Representative sections from two areas of the pancreas. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

At necropsy the proximal duodenum is often found to be hyperaemic. It is possible that this is the site of agonal vascular dilatation in bears. (P85.1.w4)

• Examination of the oesophageal musculature may detect the presence of Gonglyonema pulchrum nematodes. See: Gonglyonema pulchrum Infection in Bears
• A variety of gastro-intestinal helminths are found in bears. See:
  • Acanthocephalan Infection in Bears
  • Baylisascaris Infection in Bears
  • Cestode Infections in Bears
  • Intestinal Fluke Infection in Bears
  • Hookworm Infection in Bears
  • Gastro-Intestinal Nematode Infection in Bears

Gastrointestinal lesions may be noted with:

• Candidiasis in Waterfowl and Bears (with notes on Elephants)
• Hypersensitivity angiitis in a Bear
• Oleander Poisoning in Waterfowl and Bears
• Trichobezoars in Bears
• Yew Toxicity in Bears
• Neoplasia in Bears

Gastrointestinal parasitic infections include:

• Acanthocephalan Infection in Bears
• Baylisascaris Infection in Bears
• Cestode Infections in Bears
• Intestinal Coccidiosis in Hedgehogs (with notes on Bears) (might cause disease in juveniles)
• Cryptosporidiosis in Mammals (with special reference to Hedgehogs, Elephants and Bears)
• Dirofilaria ursi infection in Bears (submucosa of the oesophagus)
• Intestinal Fluke Infection in Bears
• Hookworm Infection in Bears
• Gastro-Intestinal Nematode Infection in Bears
• Gonglyonema pulchrum Infection in Bears (oesophagus musculature)

Check the gastro-intestinal contents for potentially poisonous plants:

• Oleander Poisoning in Waterfowl and Bears
• Yew Toxicity in Bears

Pancreatic disease includes:

• Pancreatic Necrosis in Bears
• Neoplasia in Bears

• Rupture of the stomach, intestines or caecum can occur post mortem. (B600.17.w17)
• If rupture occurred while the rabbit was alive, there will be signs such as haemorrhage, inflammation and peritonitis. (B600.17.w17)

Examine the gastrointestinal tract (GIT) in situ, noting the position and size of the stomach and caecum, and any ometal adhesions. (B611.4.w4, B614.5.w5)

To remove the GIT, cut the oesophagus close to where it enters the stomach, lift and remove the stomach and the whole of the intestines together

Stomach

• This should always contain some food. (B600.17.w17, J72.49.w4)
  • In kits, lack of milk in the stomach suggests agalactia of the dam. (J72.49.w4)
• Stomach distended, full of fluid/gas indicates:
  • Intestinal obstruction by hair mats, pulses (dried peas/beans), tumour, cysts, abscesses, adhesions or stenosis.
    • Acute Gastro-Intestinal Obstruction in Rabbits
    • Gastro-intestinal Foreign Bodies in Rabbits
  • Mucoid Enteropathy in Rabbits (terminal stages).
  • Pyloric obstruction (rare) (Pyloric Stenosis in Rabbits).
• Stomach contents impacted, often with mixed hair trichobezoar) indicates hypomotility (Gastric Stasis Syndrome in Rabbits). This is often secondary to pain/stress
  • Usually seen in rabbits which have died of Hepatic Lipidosis in Rabbits.
• Ulceration (Gastric Ulceration in Lagomorphs):
  • Associated with inappetance.
• Superficial mucosal haemorrhages, black stained - often in association with enteritis. (J72.49.w4)
• Note: tearing of the stomach wall should be considered as a post mortem lesion unless there is haemorrhage or inflammation around the tear. (J72.49.w4)
• Nematodes such as Graphidium strigosum may be found in the stomach (Gastric Nematode Infection in Lagomorphs) in domestic rabbits given access to grass. (J72.49.w4)

Small intestines

• Inflammation or haemorrhage of the intestinal wall:
  • Secondary to obstruction
  • Intestinal Coccidiosis in Hedgehogs and Lagomorphs
  • Bacterial infection.
    • Colibacillosis
  • Enterotoxaemia (note: more often affects the large intestines)
    • Clostridial Enteritis and Enterotoxicosis in Rabbits
• Distal ileum haemorrhage and necrosis:
  • Tyzzer's Disease in Lagomorphs (Clostridium piliforme infection)
• Stenosis:
  • Iatrogenic - surgery-associated.
  • Neoplasia such as lymphoma
    • Multicentric Lymphoma in Rabbits
    • Naturally Occurring Neoplasia in Lagomorphs
  • Tyzzer's Disease in Lagomorphs

Caecum and Colon

• Contents should be soft, not fluid or impacted. (B600.17.w17, J72.49.w4)
  • Bacterial infection, coccidiosis or enterotoxaemia may be suspected if the caecal contents are liquid. (J72.49.w4)
  • Samples of contents can be collected into sterile containers for detection of iota toxins (with Clostridial Enteritis and Enterotoxicosis in Rabbits) and for culture and examination for coccidia. (J72.49.w4)
• Serosal ecchymoses, with contents liquid and gassy:
  • Enterotoxaemia
    • A Gram-stained smear of contents may reveal clostridial organisms. See: Clostridial Enteritis and Enterotoxicosis in Rabbits
    • Caecal wall scrapings can be taken and examined cytologically for the presence of U-shaped or spiral Clostridium spiriforme organisms. (J72.49.w4) - caecal contents may be scanty and mucoid or bloody. (J72.49.w4)
• Gut wall oedematous, caecal contents dry and impacted:
  • Mucoid Enteropathy in Rabbits
  • Leporine dysautonomia
  • Caecal Impaction
• Caecum distended with light brown watery contents.
  • Bacterial or viral enteritis, e.g. Colibacillosis, Seen in rabbit colonies. (B600.17.w17)
• Serosal echymoses and fibrinous exudate on serosal surface:
  • Tyzzer's Disease in Lagomorphs
• Necrotic foci:
  • Hepatic Coccidiosis in Rabbits
  • Colibacillosis (in young rabbits). Clostridial Enteritis and Enterotoxicosis in Rabbits
  • Salmonellosis - particularly necrotic foci in the sacullus rotundus. (J72.49.w4)
  • Yersiniosis
  • Tyzzer's Disease in Lagomorphs
  • Tularemia
  • Listeriosis
• Note: large numbers of Passulurus ambiguus may be found (Pinworm Infection in Lagomorphs); these are not generally considered to be pathogenic. (J72.49.w4)
• In the distal colon, well-formed faecal balls should be the only contents. (J72.49.w4)
  • If mucus or excess fluid is presnet, samples should be taken for bacterial investigation. (J72.49.w4)

Mesentry

• Multiple oval cysts:
  • Cysticercus pisiformis (Tapeworm Cyst Infection in Lagomorphs)

Further investigations

• Take impression smears of the gut mucosa for cytological examination. (J72.49.w4)
• Samples of stomach and intestinal contents should be taken to look for toxins including unusual plant materials, and heavy metals. (B600.17.w17)
  • These may be seen more easily if the contents are placed in a bowl of water. (B600.17.w17)

For histopathology

• Take a 1-2 cm long piece of the relevant part of the intestine. Mark a wooden tongue depressor in pencil, indicating tissue orientation. Open the intestinal section longitudinally and pin to the tongue depressor (e.g. with 23 g hypodermic needles), serosal surface to the wood. Float this, tissue side down, in 10% formal saline in a closed container. After 24 hours or more, remove the needles and transfer the tissue to a plastic bag for labelling, sealing and proper packaging for postage. (B600.17.w17)

For anaerobic culture

• Tie off a section of intestine/caecum at both ends. Cut this section out of the intestines so that the section removes remains closed. Submit to the laboratory for culture. (B600.17.w17)
• Or take swabs of intestinal contents and push well into transport media to keep anaerobic conditions. (B600.17.w17)

For parasites

• Impression smears or intestinal scrapings can be examined for coccidial oocysts. (B600.17.w17)
• To detect worm eggs or protozoal oocysts in intestinal contents or faeces:
  • Mix the gut content/faecal sample with saturated salt solution.
  • Centrifuge.
  • Fill the container to the brim with the salt soluion.
  • Place a cover slip over the meniscus of the liquid and leave for a few minutes.
  • Remove the cover slip and place on a microscope slide.
  • Examine under the microscope.

  (B600.17.w17)

• Cut along the oesophagus and examine its mucosal surface. (B232.16.w16)
• Note whether the stomach and intestines are empty, or are full of ingesta, fluid, or gas. (B232.16.w16)
• Note any areas of inflammation on the serosal surface. (B232.16.w16)
• Open the stomach and intestines; examine the mucosal surface for lesions. (B232.16.w16)
• Note: There is no obvious junction between the small intestines and the large intestines. (B232.16.w16)
• Take a sample of intestines for bacterial culture. (B232.16.w16)

Consider:

• Atypical Mycobacterial Infection in Ferrets

• Avian Tuberculosis in Waterfowl, Lagomorphs and Ferrets (with notes on Hedgehogs)

• Campylobacter Infection in Waterfowl and Ferrets

• Colibacillosis (with special reference to Waterfowl, Hedgehogs, Elephants and Lagomorphs, and notes on Bears)

• Cryptococcosis in Ferrets

• Cryptosporidiosis in Mammals (with special reference to Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)

• Eosinophilic Gastroenteritis in Ferrets

• Epizootic Catarrhal Enteritis of Ferrets

• Gastric Ulceration in Lagomorphs and Ferrets

• Gastric Bloat Syndrome in Ferrets

• Gastro-intestinal Foreign Bodies in Rabbits and Ferrets

• Gastro-Intestinal Nematode Infection in Ferrets

• Giardia Infection in Mammals (with special reference to Hedgehogs, Lagomorphs and Ferrets)

• Inflammatory Bowel Disease in Ferrets

• Intestinal Coccidiosis in Hedgehogs, Lagomorphs and Ferrets (with notes on Bears)

• Helicobacter mustelae Gastritis in Ferrets

• Megaoesophagus in Ferrets

• Neoplasia in Ferrets

• Proliferative Bowel Disease in Ferrets

• Salmonellosis (with special reference to Waterfowl, Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)

• Rotavirus Infection in Rabbits and Ferrets with a note on Bears

• Tapeworm Infection in Ferrets

• Examine the stomach, duodenum, jejunum, ileum, caecum, appendix, colon, rectum.
• Take samples of: gastric cardia, gastric fundus & pylorus, duodenum (at the level of the bile duct, and including attached pancreas), jejunum (anterior, middle, distal), ileum, ileocaecocolic junction (and attached lymph nodes), caecum, appendix, colon (ascending, transverse, descending). For each section of intestine, open to expose the mucosa, label a piece of paper and allow the serosal surface to adhere to the paper for a moment before placing paper plus sample into formalin. Alternatively, closed pieces of bowel can have formalin gently injected into them before they are placed into the pot of formalin.
• After samples have been taken for histology, the mucosa can be washed to remove contents, or scraped, for further evaluation.
• Take samples of parasites and faeces for parasitological examination.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Superficial and deep lymph nodes from both sides of the body should be examined visually, by palpation and on cut serial section for evidence of enlargement, focal lesions, abscesses, tumour metastases, granulomata etc. 
• Weigh and record organ dimensions (maximal length and width) as required.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases.(J1.32.w7)

(B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Neoplastic lesions may be seen. See: Neoplasia in Hedgehogs

• Always consider Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs and Elephants) (D292)
  • Carefully check the tonsillar area and submandibular lymph nodes for lesions which may appear tuberculous; also check all other lymph nodes, in all parts of the body, for example the reproductive and gastro-intestinal tracts. (D292)
  • Any lymph node which appears caseous or granulomatous should be divided and (a) fixed in formalin; (b) frozen or ultrafrozen to be sent for culture. (D292)
  • Once the lungs and trachea have been removed from the thorax, at the junction of the trachea and bronchi, identify the bronchial lymph nodes, divide as above and send for histopathology and culture. This should be done even if there is no indication of TB. (D292)
  • See: D292 - Elephant Necropsy Protocol (Elephas maximus and Loxodonta africana)

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Transverse sections of the cervical, anterior, mediastinal, bronchial, mesenteric and lumbar lymph nodes. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D 293 - Elephant Research And Tissue Request Protocol

• Anthrax (with special reference to Waterfowl, Elephants and Bears) (lymph node enlargement and congestion)
• Brucellosis (experimental infection; enlarged axillary lymph nodes)
• Elokomin Fluke Fever in Bears (enlarged, soft, mesenteric lymph nodes)
• Hypersensitivity angiitis in a Bear
• Pseudorabies in Bears (Viral Disease Link) (congested mesenteric lymph nodes)
• Pythiosis in Bears (axillary and popliteal lymph nodes draining affected feet)
• Infectious Canine Hepatitis (with special reference to Bears) (mesenteric lymph nodes may be pale, enlarged, and may be petechiated.)
• Intestinal Volvulus in Bears (congested mesenteric lymph nodes)
• Salmon Poisoning in Bears (enlarged mesenteric lymph nodes)
• Neoplasia in Bears
• Hepatic and Bile Duct Neoplasia in Bears

• The superficial lymph nodes should be palpated as part of the general external examination. (B611.4.w4)

Mesenteric lymph nodes

• Oedematous, enlarged:
  • Bacterial or viral enteritis, e.g. Colibacillosis seen in rabbit colonies. (B600.17.w17, J72.49.w4)
    • In association with caecal distension with light brown watery contents. (B600.17.w17)
• Mesenteric and cervical lymph nodes oedematous with Listeriosis. (J72.49.w4)

Further Investigations

• In rabbits with Hepatic Coccidiosis in Rabbits, impression smears from the mesenteric lymph node may reveal coccidial stages of the Eimeria. (B600.10.w10, J72.49.w4)
  • Carefully cut the lymph node in half.
  • Apply the cut side to a clean microscope firmly, several times.
  • Air-dry the slide.
  • Stain the slide.
  • Examine under the microscope for the presence of sporozoites.
  • (B600.17.w17)
  (B600.17.w17)

• Check the superficial lymph nodes. (B232.16.w16)
  • Enlargement of the lymph nodes may indicate Lymphoma in Ferrets. (J29.6.w1)
  • Enlargement of a submandibular lymph node may indicate infection of the root of a canine tooth on the same side (Dental Abscesses in Ferrets). (B602.2.w2)
  • Multiple enlarged peripheral and visceral lymph nodes were seen in ferrets with Cryptococcosis in Ferrets
• Examine the mesenteric lymph nodes. (B232.16.w16)
  • Note: normal mesenteric lymph nodes are usually less than 7 mm diameter and are buried in adipose tissue. (J513.4.w5)
  • Mesenteric lymph node enlargement may be noted with Lymphoma in Ferrets and with other Neoplasia in Ferrets
  • Mesenteric lymph node enlargement and lesions are common in ferrets with Mammalian Tuberculosis (lesions also may be found in the liver, spleen, peritoneum and brain). (J195.11.w2)
    • Mesenteric lymph node enlargement has also been seen with Avian Tuberculosis in Waterfowl, Lagomorphs and Ferrets (with notes on Hedgehogs)
  • Mesenteric lymphadenopathy, hepatomegaly and splenomegaly may be present with Aleutian Disease in Ferrets. B627.15.w15, J213.8.w3)
  • Lymph nodes tend to be 8 - 12 mm, red, soft, and often oedematous with Inflammatory Bowel Disease in Ferrets. (J513.4.w5)
  • Enlarged lymph nodes may be present with Eosinophilic Gastroenteritis in Ferrets
  • With Ferret Systemic Coronavirus Infection, mesenteric lymph nodes  may be greatly enlarged (up to eight times normal), with the capsular surface irregular due to multiple slightly raised nodules, replacing the normal parenchyma. (J213.13.w1)
  • Necrotic foci may be found in the mesenteric lymph nodes of ferrets with Salmonellosis. (B627.14.w14)

• Take sections of axillary and inguinal lymph nodes (section transversely through the hilus).
• Take sections of mandibular lymph nodes with the mandibular salivary glands.
• Section cervical lymph nodes transversely.
• Section mesenteric lymph nodes transversely. Include colonic lymph nodes with sections of colon.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Autolysis of the brain occurs particularly rapidly. Priority should be given to removal of the brain for fixation if required.
• In areas where rabies infection (See: Rabies) is enzootic, all mammals found dead, and particularly those with a clinical history of abnormal behaviour or neurological signs, should be carefully examined and considered as potentially infected until proven otherwise. (See: Human Health Considerations section on this page) 
  • Specialist examination of the brain is required for diagnosis of rabies infection.
• Removal of the brain and spinal cord may require experience and specialist equipment, particularly for dealing with very large or small taxa, and those with particularly robust skulls (e.g. Suidae - Pigs (Family))
• First disarticulate the skull at the level of the atlanto-occipital joint and remove all overlying musculature.
• Use an appropriate saw to cut through the cranium; techniques may vary with the species under examination.
  • Make a transverse cut across the width of the skull just behind the orbits (eyes).
  • Make two cuts joining with the outer limits of the first cut, towards the foramen magnum.
  • Alternatively cut around the skull cap.
  • A medical vice used to stabilise the skull in position may be useful if available.
• Carefully lift off the top of the skull using blunt elevation.
• Examine the brain and meninges in situ for evidence of local haemorrhage, bruising, swelling, redness or pale colouration, parasites, abscesses, growths or inflammation, etc.
• Remove the brain from the skull through careful elevation, cutting the cranial nerves in turn.
• Weigh and record organ dimensions (maximal length and width) as required.
• Examine the cut surface of serial sections of the brain, noting the distribution of any lesions found.
  • The distribution of focal lesions found on post mortem examination should be related to the clinical history where neurological signs were noted. 
• Dissection for examination of the spinal cord can be time consuming.
  • Dissect away the overlying musculature, carefully trim the vertebral arches to allow visualisation of the spinal cord.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Fixation of the entire brain may be required. In this instance, it may be helpful to remove the top of the skull and place the skull and brain in fixative overnight. The brain will then become increasingly firm and can be easily removed from the skull without damage.
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Samples for toxicology should be either wrapped in foil ( if the suspected toxin is organic), plastic wrap ( if the suspected toxin is metal) or glass (if suspected toxin is unknown).
  • Brain samples are important for toxicological investigation (e.g. organochlorines, organophosphates, carbamates).
• Appropriate chain of custody and sample labeling should be followed when dealing with legal cases.(J1.32.w7)

(B273, B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• The opening of the cranium is best conducted after separating the head from the body. A chainsaw, large axe and chisels are needed for the procedure. (B450.B.w28)

• Any abnormalities affecting the brain, meninges, spinal cord and peripheral nerves. (B450.B.w28)

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Representative sections of the brain. (B450.B.w28)
• A three cm section of the sciatic nerve. (B450.B.w28)

Requested measurements to be taken and tissues/features to be looked for, as well as information on samples to be taken for TB and EEHV are provided in: D293 - Elephant Research And Tissue Request Protocol

• An unusual fluke infection described in one bear included involvement of the CNS. See: Prouterina wescotti Infection in Bears
• Toxoplasmosis in Waterfowl (with notes on Hedgehogs, Elephants and Bears) has been seen causing CNS lesions in bears.
• Gross changes with enlargement of the ventricles is seen with Hydrocephalus in Bears
• CNS lesions including extensive occlusive fibrinopurulent exudate within the cerebral ventricles was noted in a cub with Candidiasis in Waterfowl and Bears (with notes on Elephants)
• No lesions may be found associated with Epilepsy and Convulsions in Bears
• Other diseases with CNS lesions include:
  • Brain Infarct in Bears
  • Cerebral Arteriosclerosis in a Bear
  • Cretinism in a Bear
  • Neuronal Degeneration of the Spinal Cord in Bears
  • Ruptured Intervertebral Disk in a Bear

Brain

• To remove from the skull, carefully remove a section of thin parietal bone at the base of the skull.
• Make a midline incision in the sulcus so formal saline can penetrate the ventricles.
• Fix the whole head.
  • Or (harder to keep the brain intact) carefully dissect the brain from the surrounding tissues, remove and fix. (B600.17.w17)
• Histopathological examination can be used to confirm Encephalitozoonosis in Lagomorphs (J72.49.w4)

Note: If the whole head is fixed and then the bones decalcified (in decalcifying solution) serial sections can be taken to show lesions such as abscesses in the nasal passages, tympanic bullae or cranium, as well as tooth roots and their relationships to surrounding tissues. (B600.17.w17)

Spinal cord

• Radiography can be used to visualise lesions. (B600.17.w17)
• To remove the spinal cord for further examination: (B600.17.w17)
  • Dissect away the muscles to reveal the vertebral column.
  • Using nail clippers, gently remove the laminae.
  • Transect the spinal cord (proximal and distal) and spinal nerves.
  • Lift the spinal cord out of the vertebral canal.
  • Inspect the floor of the canal for compressive or traumatic lesions.
  • Note: Fixing the tissues first makes it easier to remove the spinal cord intact. (B600.17.w17)

Autonomic ganglia

• These are examined if Leporine dysautonomia is suspected, in lagomorphs which have dies with intestinal motility disorders (particularly constipative mucoid enteropathy). (B600.17.w17)
• The cranial mesenteric, caudal mesenteric, stellate gangla and cranial ganglia, as well as the brain, can be assessed for neuropathological changes seen in dysautonomia. (B600.17.w17)
• Note: it is important to take care in locating and removing the ganglia to avoid fragmenting or crushing them. They should be placed in a container separate from other tissues intended for histopathological examination. (B600.17.w17)
• To remove the autonomic ganglia: (B600.17.w17)
  • Place the rabbit on its right side.
  • Remove the left abdominal and thoracic walls.
  • For all the ganglia, use the flat edge of a No. 22 scalpal blade to remove the soft fat overlying the ganglia and reveal the firmer, semitransparent ganglia.
  • To find the cranial mesenteric ganglion, look on the left side of the aorta, ventro-cranial to the left adrenal.
    • If this is dissected out properly, with care not to crush it or fragment it, it can provide enough material for diagnosis even if other ganglia are not obtained. 
  • To find the caudal mesenteric ganglion (very small), look in the roof of the mesentry subtending the distal colon.
    • The ganglion is long and thin. Take care to distinguish it from the lymph nodes (more friable) beneath it.
  • To find the left stellate ganglion, transect the trachea and oesophagus in the middle of the trachea. Deflect these , exposing the tissues at the entrance to the right side of the chest, beneath the first rib.
  • To find the cranial cervical ganglion, follow the vagosympathetic trunk cranially from the midneck to the base of the skull. The ganglion is a tiny fusiform swelling in the proximal end of the sympathetic trunk, close to the nodose ganglion (slightly larger) on the adjacent vagus.

  (B600.17.w17)

• If there is any concern regarding the possibility of Rabies, the head must be sent to an appropriate laboratory for examination of the brain.

  • In areas where rabies occurs, consider rabies in any ferret with paralysis or sudden behavioural/personality change, particularly if they have access to outside, a rabies epizootic is occurring locally, or the ferret is unvaccinated. (J213.8.w3)

• Fix the brain either whole or after sectioning transversely.
• Allow the brain to fix for at least one week before sectioning transverselly into slabs 0.5 - 1.0 cm thick to examine for lesions.
• Either send the whole brain to the pathologist for histopathological examination or transverse slabs should be submitted from the medulla, pons, cerebellum, midbrain, thalamus and cortex (prefrontal, parietal, occipital including hippocampus and lateral ventricles with choroid plexus).
• Fix the whole pituitary.
• Remove and fix the trigeminal ganglion (Gasserian ganglion).

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment

• Incise and examine the major limb joints in turn, comparing both limbs for reference in each case.
  • Note the volume, colour, viscosity and turbidity of synovial fluid present.
  • Examine joint surfaces and tendon sheaths for evidence of discolouration, irregularity, degeneration, joint mice, etc. 
• Check the state of mineralisation of the major limb bones by seeing how difficult they are to bend or break.
• Incise and examine the leg muscles over multiple areas of the body.
  • Note the muscle colour (e.g. pale areas with white muscle disease and capture myopathy (See: Capture Myopathy), dark appearance with clostridial infection), any haemorrhage present. 
• Examination of the bone marrow can be used during the post mortem examination of ungulates as a judge of nutritional plane; animals on a healthy, adequate diet will have yellow/ white firm and fatty marrow in comparison with orange/red, watery and soft marrow in animals suffering from chronic starvation or debility.

Samples

• Take tissue/swab for culture, impression smears from cut surface, tissue for histopathology and toxicology as appropriate.
• Tissue to be fixed for histopathology should be no more than 10mm thick, and thinner if congested. 
• Samples for histopathology should not be frozen since this results in crystal formation and cell rupture.
• Samples for histopathology should be placed in at least ten times the volume of formalin fixative as the tissue.
• Appropriate chain of custody and sample labelling should be followed when dealing with legal cases.(J1.32.w7)

(B273, B411.II.w2, B411.III.w3, J1.32.w7, P24.327.w13, D91.4.w1, V.w26)

• Check carefully for limb fractures in elephants which have collapsed in an awkward position. (B411.IV.w4, P80.1.w1)
• All parts of the fore and hind limbs should be examined. This requires the limbs to be disarticulated at the major joints (shoulder, elbow, radio-ulnar and carpal joints of the fore-limbs, hip, stifle, tibiofibular and tarsal joints or the hind limbs). (D286.2.w2)
  • A small axe my be used to cut through the joints; a crowbar may be needed to lift parts of the leg and for separating the joints. (D286.2.w2)
• "Note any pathological conditions in the joints. Slight erosions on articular surfaces can be viewed best in fresh tissues and should be examined soon after death. Grooves and fractures on articular surfaces cannot be mistaken and should be sought. Look also for "joint mice," calcium deposits, and any other abnormal signs." (D293)
• Incise through the muscles in multiple locations. (D286.2.w2)
• Check for lesions of capture myopathy. (P80.1.w1)
• See: 

  • Bone Fractures in Elephants

  • Calcium-Vitamin D Metabolism Imbalance (with special reference to Hedgehogs and Elephants)

  • Capture Myopathy (with special reference to Waterfowl and notes on Elephants)

  • Cardiac Glycoside Toxicity in Elephants

  • Clostridium septicum Infection in Elephants

  • Degenerative Joint Disease in Elephants

  • Encephalomyocarditis (with special reference to Elephants)

  • Flaccid Trunk Paralysis in African Elephants Loxodonta africana

  • Foot-and-Mouth Disease
    • (Note: Details of the disease in elephants are given on the page Foot and Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA)

  • Foot Wounds in Elephants

  • Joint Luxations in Elephants

  • Mycoplasma Infection (with special reference to Waterfowl, Hedgehogs and Elephants)

  • Rabies (with special reference to Waterfowl, Hedgehogs and Elephants)

  • Sunstroke - Heatstroke in Waterfowl and Elephants)

  • Suprascapular paralysis in Elephants

  • Tetanus (with special reference to Elephants)

  • Traumatic Musculoskeletal Swellings in Elephants

  • Trunk Paralysis in Asian Elephants Elephas maximus

  • Tutu (Coriaria arborea) Toxicity in Elephants

  • Varagu (Paspalum serobiculatum) Toxicity in Elephants

  • Vitamin E - Selenium Deficiency (with special reference to Waterfowl and Elephants)

Recommended tissue sampling: (B450.B.w28)

• Gross lesions. 
• Section of the rib, including the costochondral junction. (B450.B.w28)
• Skeletal muscle, a cross section of the thigh muscle. (B450.B.w28)

• Arthritis and Skeletal Disease in Bears
• Calcium-Vitamin D Metabolism Imbalance (with special reference to Hedgehogs, Elephants and Bears)
• Clostridial Myonecrosis in Bears
• Fractures in Bears
• Lacerations & Punctures, including bite wounds (with special reference to Waterfowl, UK Wildlife and Bears)
• Radial Agenesis in a Bear
• Ruptured Intervertebral Disk in a Bear

Various parasites may be found in the muscles of bears. See:

• Cysticercosis in Bears
• Sarcocystosis in Waterfowl and Bears
• Sparganosis in Bears
• Toxoplasmosis in Waterfowl (with notes on Hedgehogs, Elephants and Bears)
• Trichinella Infection in Hedgehogs and Bears

The bone marrow may be non-fatty in bears which have been starving. See: 

• Starvation in Waterfowl and Seabirds (with notes on Hedgehogs and Bears)

• The muscles of the rabbit are normally quite pale, not dissimilar to those in the domestic chicken. (J72.49.w4)
• Palpate for fractures. (J72.49.w4)
• Examination for fracture or dislocation of lumbar vertebrae may be carried out most easily by taking a whole-body radiograph. (J72.49.w4) See: Imaging in Lagomorph Diagnosis and Treatment - Radiography
• Abnormalities which may be detected include:
  • Fractures in Rabbits
  • Elbow Luxation in Rabbits
  • Hip Dislocation in Rabbits
  • Intervertebral Disc Disease in Rabbits
  • Spina Bifida in Rabbits (in neonates)
  • Spinal Abnormalities in Rabbits
  • Spinal Injury in Lagomorphs
  • Splay Leg in Lagomorphs (in neonates or juveniles)
  • Traumatic Jaw Injuries in Lagomorphs

Check for lesions of:

• Fractures in Ferrets
• Intervertebral Disc Disease in Rabbits and Ferrets
• Neoplasia in Ferrets (particularly chordomas). (B602.9.w9)
• Spinal Abscess in Ferrets
• Spinal Injury in Lagomorphs and Ferrets

• Examine the muscles.
• Examine the joints.
• Open the femur to check the bone marrow.
• Examine the spinal column, dorsal and ventral aspects.
• Fix complete sections or hemisections of bone marrow from the femur.

(D408)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment
• Imaging in Lagomorph Diagnosis and Treatment - Radiography

• Potential hazards to human health of ANY necropsy or post mortem examination must be considered before undertaking examinations.
• Personnel undertaking or attending necropsies or post mortem examinations must be made aware of the potential hazards to human health.
• All post mortem examinations should be considered to be harbouring potentially transmissible agents of infectious disease until proven otherwise and should be handled accordingly with due caution.
• Potential hazards range from toxins on the surface of the animal (e.g. oil) to zoonotic diseases which may be transmitted through cuts, absorbed through mucous membranes or inhaled in the form of dust or aerosols.
• Important zoonotic diseases to consider when dealing with mammal carcasses include, amongst others, Salmonellosis, Rabies, Anthrax, Echinococcus spp. (Cestoda - Tapeworms), Mammalian Tuberculosis, Brucellosis.
  • Zoonoses associated with particular taxa include Herpes simiae (Herpes B) (Herpesviridae) in macaques (Cercopithecidae - Old-world monkeys (Family)); Hendra virus and Australian bat lyssavirus (Rhabdoviridae) in bats (Chiroptera - Bats (Order)); tulareamia, plague (Yersinia) and hantavirus (Bunyaviridae) in rodents (Rodentia - Rodents (Order).
  • Zoonoses where knowledge of the natural wildlife host is incomplete include Ebola Virus (Filoviridae) in Central Africa and Nipah virus in Malaysia.
• For all necropsies, protective clothing should be worn including:
  • Disposable gloves (which should be replaced immediately if damaged). 
  • Overalls (material suitable for boil clean).
  • Washable apron.
  • Rubber boots.
  • A face mask, which covers the nose and mouth, is advisable, particularly when aerosol transmission infections are suspected. 
  • Protective eye wear may be recommended to prevent ocular splash injuries.
• If possible, necropsy should be carried out inside a protective cabinet. This is highly recommended when examining primate carcasses.
• Any cuts should be washed immediately with a disinfectant soap.
• Appropriate regional health and safety legislation should be consulted as necessary.

(B273, B411.I.w1, D91.4.w1, V.w26)

• Zoonotic infections reported to occur in the hedgehog include:
  • Bordetella bronchiseptica Infection [Infections in humans are usually "under atypical conditions" (P23.199S.w8)]
  • Chiggers Infection
  • Colibacillosis [Particularly vero cytotoxic Escherichia coli serogroup 0157]
  • Cryptosporidiosis [Very wide host range (P23.199S.w8)]
  • Erysipelothrix Infection [Via bites or skin punctures caused by spines (P23.199S.w8)]
  • Flea Infection [Fleas may jump onto humans; some people appear more attractive to fleas than others (P23.199S.w8)]
  • Foot-and-Mouth Disease [Rarely infects humans]
  • Giardia Infection [Very wide host range (P23.199S.w8)]
  • Leptospirosis [Risk from urine-contaminated food or water - oral route of infection or skin contamination (P23.199S.w8)]
  • Louping-ill [Theoretically could be present in sheep ticks Ixodes ricinus which are found on hedgehogs (P23.199S.w8)]
  • Lungworm Infection [Capillaria aerophila infection may rarely occur in humans (P23.199S.w8)]
  • Lyme Disease [Transmitted by ticks (P23.199S.w8)]
  • Mammalian Tuberculosis [Not recorded to date in hedgehogs in the UK, but has been isolated from hedgehogs in New Zealand]
  • Pasteurella Infection [Via bites or skin punctures caused by spines (P23.199S.w8)]
  • Pseudomonas Infection [Via bites or skin punctures caused by spines (P23.199S.w8)]
  • Q Fever [Transmitted by ticks]
  • Rabies [Not present in the UK at this time (2002). Rare in hedgehogs but self-anointing, involving profuse salivation, may lead to suspicion: see West European hedgehog Erinaceus europaeus - Activity Patterns, Grooming and Navigation Behaviour (Literature Reports)]
  • Ringworm [There are a number of records of human ringworm caught from hedgehogs]
  • Salmonellosis [There are a number of records of human salmonellosis associated with salmonellosis in hedgehogs]
  • Staphylococcosis [Via bites or skin punctures caused by spines (P23.199S.w8)]
  • Sarcoptic mange
  • Tick Infection
  • Toxoplasmosis [Unlikely to be a problem unless the hedgehog is eaten without proper cooking (V.w5)]
  • Trichinella Infection  [Unlikely to be a problem unless the hedgehog is eaten without proper cooking (V.w5)]
  • Yersiniosis [Yersinia enterocolitica, which has net been found in hedgehogs, is more likely to cause disease in humans than is Yersinia pseudotuberculosis. (B209.19.w19, P23.1999S.w8)]

Documented zoonotic transmission in elephants include:

• Anthrax
• Poxvirus Infection in Elephants
• Mammalian Tuberculosis

Potential zoonoses:

• Cryptosporidiosis in Mammals
• Encephalomyocarditis
• Foot-and-Mouth Disease
  • (Note: Details of the disease in elephants are given on the page Foot and Mouth Disease - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA)
• Leptospirosis
• Rabies
• Ringworm
• Salmonellosis
• Mycoplasma Infection
• Toxoplasmosis
• Yersiniosis

• Anthrax (with special reference to Waterfowl, Elephants and Bears) [Very rare; two cases reported]
• Baylisascaris Infection in Bears [Larvae of the raccoon roundworm Baylisascaris procyonis can infect humans; it is not known whether larvae of Baylisascaris transfuga can infect humans]
• Chiggers (Trombiculiasis) (In Waterfowl, Hedgehogs and Bears)
• Colibacillosis (with special reference to Waterfowl, Hedgehogs and Elephants and a note on Bears)
• Rabies (with special reference to raccoons) (Viral Disease)
  • Information regarding the disease in bears is available in:
    • Rabies (with special reference to Raccoons) - Detailed Clinical Signs (Disease Reports)
    • Rabies (with special reference to Raccoons) - Detailed Pathological Findings - (Necropsy-Post Mortem) (Disease Reports)
    • Rabies virus (with special reference to raccoon rabies variant) - Definitive Mammal Host Species (Viral Reports)
    • Rabies virus (with special reference to raccoon rabies variant) - Detection and Identification Techniques (Viral Reports)
• Ringworm in Hedgehogs (with notes on Elephants and Bears)
• Salmonellosis (with special reference to Waterfowl, Hedgehogs, Elephants and Bears)
• Sarcoptic Mange
• Tick Infection (with special reference to Hedgehogs, Elephants and Bears)
• Toxoplasmosis in Waterfowl (with notes on Hedgehogs, Elephants and Bears) [Zoonotic potential only if raw or undercooked bear meat is eaten]
• Trichinella Infection in Hedgehogs and Bears [Zoonotic potential only if raw or undercooked bear meat is eaten]
• Yersiniosis (with special reference to Waterfowl, Hedgehogs, Bears and notes on Elephants) (Bacterial Disease) [Rare zoonosis; can be severe or fatal]

Domestic rabbits

• There are negligible risks of transmission of serious zoonotic diseases from pet rabbits to healthy humans. (B600.3.w3)
• Appropriate protective clothing should be worn; when zoonotic disease is suspected, particularly if the handler is immunocompromised, extra hygienic precautions should be taken. (B601.2.w2)
• Allergy to rabbit dander can develop. (B600.3.w3)
• Parasites which may be transmitted from rabbits to humans include:
  • Fleas (Flea Infection in Mammals) (usually cat or dog fleas rather than rabbit fleas). (B600.3.w3)
  • Cheyletiella parasitivorax (see Cheyletiellosis). (B600.3.w3)
    • In humans, this results in pruritic, erythematous lesions, usually found on the arms. (B600.3.w3)
  • Toxoplasma gondii (see Toxoplasmosis) (but only by eating undercooked rabbit meat, not by handling a rabbit or its faeces). (B600.3.w3)
  • Encephalitozoon cuniculi (Encephalitozoonosis in Lagomorphs) (only in immunocompromised people). (B600.3.w3)
• Note: while some protozoa such as Giardia can infect both rabbits and humans, there does not seem to be transmission from one species to the other. (B600.3.w3)

Wild lagomorphs

Zoonotic diseases which are known to occur in wild lagomorphs include:

• Tularemia (B600.3.w3)
• Plague (Yersinia pestis infection - Yersiniosis). (B600.3.w3)
• Listeriosis. (B600.3.w3)
• Rabies. [Not in the UK. Lagomorphs are not major hosts of this disease]

Viral

• Influenza in Ferrets This may be passed from human to ferret and from ferret to human. (P120.2007.w1)
• Rabies

  • See: Rabies - Editorial Overview of Disease Characteristics for specified SPECIES-TAXA

  • In areas where rabies occurs, consider rabies in any ferret with paralysis or sudden behavioural/personality change, particularly if they have access to outside, a rabies epizootic is occurring locally, or the ferret is unvaccinated. (J213.8.w3)

  • Rabies is not common in ferrets; during 1960 - 2000 in the US, 2,851 cases of rabies were diagnosed in carnivores other than raccoons, skunks (Mephitis, Spilogale and Putorius (Mustelidae - Weasels (Family))) and foxes (Vulpes, Urocyon and Alopex). Of these cases, only 23 were in ferrets. (J1.39.w3)

  • Ferrets with rabies often do not shed virus, but shedding in the saliva may vary depending on the rabies variant; shedding was more common in ferrets infected with the raccoon rabies variant. (B627.15.w15, J13.59.w1)

• Campylobacter Infection in Waterfowl and Ferrets
• Leptospirosis (with notes on Hedgehogs, Elephants, Bears and Lagomorphs)
• Listeriosis in Waterfowl and Lagomorphs (with notes on Hedgehogs, Bears and Ferrets)
• Salmonellosis (with special reference to Waterfowl, Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)
• Mammalian Tuberculosis (with special reference to Badgers, Hedgehogs, Elephants, Bears, Lagomorphs and Ferrets)

• Ringworm

• Ectoparasites
  • Flea Infection in Mammals
  • Otodectes cynotis Ear Mite Infection in Hedgehogs and Ferrets
  • Sarcoptic Mange
• Endoparasites:
  • Gastro-Intestinal Nematode Infection in Ferrets
  • Heartworm Infection in Bears and Ferrets (with a note on Lagomorphs)
  • Tapeworm Infection in Ferrets [via fleas]
• Protozoa
  • Cryptosporidiosis in Mammals
  • Giardia Infection in Mammals (with special reference to Hedgehogs, Lagomorphs and Ferrets)
  • Toxoplasmosis [but only a zoonotic risk if you eat the ferret!]

• Many diseases of great apes are zoonotic.
• Any primate that dies during quarantine should be assumed to be of high zoonotic potential until proven otherwise. (D428.8.2.w8b)
• Appropriate precautions should be taken such as wearing of appropriate personal protective equipment, including Wellington boots, aprons and other waterproof clothing, double-gloving and wearing an anti-cut glove on the non-cutting hand. (V.w150)
• When conducting a necropsy on a primate which is too large to fit into a safety cabinet, it is recommended that a powered respirator with full face visor and hood should be worn. (V.w150)
• After the necropsy, instruments, surfaces and the flood should be flushed with water then washed with a lipolytic detergent, then disinfected with e.g. diluted bleach. (D410)
• Specimen containers (outside surfaces) also should be cleaned with disinfectant then placed into sealed bags for transportation to diagnostic laboratories. (D410)
• Disposable personal protective equipment should be double-bagegd: placed into a bag which is then sprayed with disinfectant before being placed inside another bag for incineration. (D410)
• Reusable personal protective clothing should be disinfected on exiting the necropsy room. (D410)
• Primate carcasses should be disposed off properly e.g. by incineration. If a primate carcass is to be donated e.g. to a museum, it is recommended that it should be double-bagged and stored frozen until results of diagnostic tests, incuding pathogen cultures, are received. (D410)

• Chapter 2 - B36 Field Manual of Wildlife Diseases - Specimen Collection and Preservation
• Chapter 3 - B36 Field Manual of Wildlife Diseases - Specimen shipment