SOP FOR PROPER COLLECTION , PRESERVATION , DESPACTCH AND HANDLING OF DIAGNOSTIC SAMPLES FOR POULTRY

SOP FOR PROPER COLLECTION , PRESERVATION , DESPACTCH AND HANDLING OF DIAGNOSTIC SAMPLES FOR POULTRY :
Compiled, & shared by-DR. RAJESH KUMAR SINGH, (LIVESTOCK & POULTRY CONSULTANT), JAMSHEDPUR Post no 1396 Dt 09/09//2019
JHARKHAND,INDIA 9431309542, rajeshsinghvet@gmail.com

The capability of a laboratory to confirm the diagnosis of a suspected infectious animal disease is directly related to the types, amount and conditions of the specimens submitted. The field diagnostician must select, aseptically procure, and properly preserve specimens for the isolation or demonstration of a causative agent. In addition, an adequate number of specimens must be taken from the appropriate tissues, at the proper stage of the disease, to maximize the chances of isolating the pathogen. The herd or flock owner, private practitioner and diagnositc laboratory comprise a front-line defense and will, most likely, be confronted with the initial case of an infectious animal disease. It is vitally important that these people contact the state Veterinarian or Veterinarian in charge district diagnostic laboratory as soon as possible if contagious disease is suspected. The diagnostician is responsible for collecting and dispatching specimens for further investigations if needed, at laboratory of Veterinary college of respective state. When the existance of an exotic disease is suspected, no animal or specimen should be removed from the premises of origin unless in the custody of an officially designated diagnostician.

SEROLOGY AND BLOOD COLLECTION————
Diagnostic samples are used to determine health status or identify specific pathogens in pullet, layer and breeder flocks. Routine samples include whole blood, serum, formalin-fixed tissue and swabs:
tracheal, choanal, oropharyngeal, cloacal, organs and joints. For specific investigations, Fast Technology for Analysis of nucleic acids (FTA) cards can be used to collect feather pulp, whole blood or isolates from any type of swab.
SAMPLE SUBMISSION —————-
When submitting samples to a diagnostic laboratory, it is important to provide thorough and relevant flock information on the laboratory submission form. Critical information that should accompany all diagnostic sample submissions includes:
• Flock identification and location
• Age of flock
• Date of sample collection
• Vaccination program
• Flock history, including pertinent health or production problems
This information is vital to the flock veterinarian and diagnostician to make a meaningful interpretation of serological or diagnostic results and provide recommendations to improve flock health and/or production.

Summary of Guidelines for Proper Serum Collection —————

• Select normal representative birds (10 to 20 sera samples), unless working up a diagnosis.
• Collect 2.0 to 3.0 mL of blood from each bird.
• Samples collected with a needle are cleaner than with a scalpel.
• Do not damage samples by forcing the blood sample back through the needle into the clot tube.
• Ensure blood runs down the side of the clot tube and position the tubes nearly flat until the clot is formed.
• Leave blood in the clot tube for 10 to 12 hours at about 80°F (27°C).
• Do not shake, roughly handle or freeze the blood while the clot is forming or hemolysis will occur.
• Remove clot gently, or pour off serum.
• Do not mail samples without first removing the clot.
• Keep the serum samples cool and send immediately to the laboratory on wet ice or cold pack.

Ages for blood collection in breeder flocks:————-

1. 10 to 12 weeks
2. 2. At time of transfer (grow to lay farm)
3. 3. Every 10 to 12 weeks during egg production
Ages for blood collection in commercial layer flocks: ——-
1. One time prior to transfer (grow to lay farm)
2. Every 10 to 12 weeks during egg production

SEROLOGY———–

Serology is the study of serum antibody levels, also known as titers. The immune system develops antibodies that circulate in the blood after a bird is exposed to an antigen, whether by vaccination or exposure to a wild-strain pathogen. Antibodies are found in the serum portion of blood (the liquid portion after the clot develops). Serum is free of all blood cells and clotting factors. The flock’s serum antibody titers are used to monitor efficacy of vaccination programs, evaluate field challenges or diagnose disease. The value of this information depends on the quality of the serum samples received by the laboratory. Poor quality samples lead to erroneous and misleading results. Selection of birds for blood collection, techniques used to collect blood, and handling of blood samples and serum all influence laboratory results.

Selection of Birds ————–

For routine serological monitoring, serum samples should be collected from normal, healthy birds. Do not use cull birds that are sick or appear distressed, as their antibody titers are not typically representative of the overall flock health status. During a potential disease investigation, however, blood samples should be collected from birds that are exhibiting the clinical signs or lesions of the suspected pathogen or syndrome. In caged housing systems, it is important to select birds from various locations throughout the house. When a flock is enrolled in a routine serology program, collecting blood from the same birds (or same cages) is recommended. This will reduce the variability of results when compared to collecting blood from different birds at each time of testing. In floor housing systems, identifying the same birds is difficult. Large plastic wing bands or feathers marked with dye can be useful to allow consistent collection.

Number of Samples ————

Twenty good quality serum samples should be collected for routine flock profiling and for disease investigation; however, a minimum of 10 samples may be sufficient to estimate flock antibody titers.

Ages for Sampling ———–

For routine monitoring, the first blood collection should be 10 to 12 weeks of age. By this age, a pullet flock has an opportunity to respond to early live vaccinations and maternal antibodies are absent. Antibody titers from this age group can be used to assess the overall immune status of a young flock and priming effect of live vaccines used in vaccination programs. This early serology assessment can screen for potential disease challenge in the grow house. Another important time for antibody titer evaluation is immediately prior to transfer of the pullet flock to the laying house. This is a good time to check the pullet’s immune response against Mycoplasma gallisepticum (MG), Mycoplasma synoviae (MS), Newcastle disease (NDV), infectious bronchitis (IB), avian encephalomyelitis (AE), and avian influenza (AI). In breeders, transfer is also an ideal time to assess adequate sero-conversion for chicken anemia virus (CAV) and avian encephalomyelitis (AE). Collecting serum before transfer establishes a baseline titer level for a flock moved to a multi-age complex. Titer response from inactivated (killed) vaccines will peak at 3 to 5 weeks post-vaccination. When monitoring flocks during the egg production period, a 10 to 12 week interval is sufficient to monitor changes in antibody titer levels. During a disease outbreak investigation, blood should be collected when clinical signs of the disease are first observed, followed by an additional blood collection from the same birds 3 to 5 weeks later. This collection time frame allows for specific antibody production against a potential disease agent. The comparison of titers from these paired sera samples may demonstrate significant changes in titers for a suspected pathogen. Reserving the first serum sample (by freezing) to be run at the same time as the second sample reduces lab test variance due to external factors or changes in reagent lots. A similar tactic can be used to monitor the efficacy of killed vaccines given in a pullet program.

Volume of the Blood Sample ————–

With proper collection and handling technique, 2.0 to 3.0 milliliters (mL or cc) of whole blood will yield 1.0 to 1.5 mL of serum. This volume of serum is sufficient for routine ELISA testing for Newcastle disease, infectious bronchitis, infectious bursal disease (IBD or Gumboro), AE and AI by agar gel immunodiffusion (AGID), as well as for MG, MS and pullorum-typhoid (PT) by plate agglutination testing. Sufficient serum should be kept frozen in reserve, in case additional testing is required in the future.

Equipment Used for Blood Collection—————

Disposable, sterile 3 or 5 cc syringes are used, depending on the size of the sample to be obtained. The size of needle depends on the anatomical site used for blood collection.

Blood Collection Site Needle Length Needle Gauge
Wing vein 0.5–1.0 inch 20–22 gauge
Cardiac puncture 1.5 inch 18–20 gauge

Always use disposable needles and replace needles every 5 to 10 birds. Dull needles cause tissue trauma and make accurate punctures of veins more difficult. All blood collection equipment must be changed between flocks to eliminate the potential for disease transmission. Rinsing the needle and syringe between birds with distilled water prevents blood from clotting within the needle. Sterile 3.0 mL plastic or glass blood tubes with leak-proof tops are ideal for blood collection and storage, as they allow for proper clotting of samples. Similar tubes are ideal for storing separated serum as well.

METHODS USED FOR COLLECTING A BLOOD SAMPLE————-

1. Wing (brachial) vein method using a needle
The brachial vein of the wing is an acceptable site for blood collection for birds 4 weeks and older. In younger birds, this vein is too small for efficient blood collection.
STEP 1
Hold bird by both legs.
STEP 2
Place legs under elbow of nondominant hand.
STEP 3
Free both hands to gain access to underside of wing.
STEP 4
Remove feathers to better view the brachial vein.
STEP 5
Visualize the brachial vein.
STEP 6
Orient needle in alignment with vein, bevel pointed up, with tip of needle pointed toward wing tip.
STEP 7
Needle should be inserted first under the skin and then into the vein mid-way between elbow and shoulder joints.
STEP 8
If needle is within the brachial vein, blood will fill syringe with minimal pull on syringe plunger. Pulling back on plunger with too much force will create high negative pressure, causing the vein to collapse and stopping the flow of blood into the needle.
STEP 9
Once the needle is removed from the vein, the application of slight pressure with a finger over the injection site will promote more rapid clotting. Formation of a hematoma or blood clot in the injection area is common.
All needles should be discarded in a designated sharps container. Needles should never be recapped.
If a hematoma forms before a sufficient quantity of blood has been obtained, it may be necessary to stop and attempt collection from the bird’s opposite brachial vein. Once a hematoma has formed, it is nearly impossible to visualize the vein and thus impossible to collect blood.
If blood is not flowing into the syringe:———
1. Needle is not in the vein.
2. 2. Needle is plugged with a clot.
3. 3. Vein has been punctured and a hematoma is forming.

4. Wing vein puncture using a scalpel blade—————–

Although this method can provide more rapid blood collection, it does have the potential to induce more trauma than using a needle and syringe.
a. A #11 scalpel blade inserted into a #3 or #4 scalpel blade holder is used to puncture the brachial vein just above the elbow joint.
b. A blood tube is used to collect the blood as it hemorrhages from the cut. This method is more likely to result in sample contamination with bacteria, mold, etc. Wiping the skin with rubbing alcohol prior to the cut may limit contamination.
c. Depending on the size of the cut, this method can cause significant trauma (blood loss, stress, etc.) to the bird and involves risk of severing the brachial artery and nerve.

5. Cardiac puncture methods ——-
6.
Collecting blood directly from the heart can provide rapid blood collection, and allow for collection of larger volumes of blood (4 to 10 mL). Additionally, cleaner blood samples can be collected compared with wing vein method. Cardiac puncture methods should only be practiced by trained personnel. Poor technique in needle placement and repeated attempts to locate the heart can result in fatal hemorrhage; however, this risk is minimized with practice. If fatal hemorrhage is suspected, the bird should be humanely euthanized promptly.

a. Anterior (thoracic) cardiac approach——-
This is a one-person technique where the bird is restrained by holding both legs in one hand while operating the syringe with the other hand. The proper position of the bird is flat on its back with the bird’s head extended downward over the edge of a table (or cage or handler’s knee). Using the index finger as a guide, the needle is inserted into the thoracic inlet at the highest point of the inverted V formed by the clavicle (wishbone). The needle is kept in the same plane as the keel bone and angled back toward the tail. The entire length of the needle (1.5 inch or 3.81 cm, 18 gauge) is usually inserted with little resistance into the heart. While inserting the needle, a slight negative pressure is applied. When the needle enters the heart, the blood will flow easily into the syringe. When the needle is positioned incorrectly, usually not in the same plane as the keel bone, it can enter the respiratory tract and air will flow back into the syringe. Hemmorhage into the lungs or airsacs can result from needles positioned incorrectly. Should incorrect needle insertion occur, and evidence of respiratory distress is observed, the bird should be humanely euthanized in an appropriate manner.

b. Lateral cardiac approach ———-
c.
A lateral approach is practiced by inserting a needle through the left thoracic wall. This is usually a two-person procedure where the bird handler lays the bird flat on a table on its right side, holding both legs in one hand and both wings in the other. The landmark is the groove formed by the edge of the breast (pectoral) muscle, where the ribs can be felt. A 1.5 inch (3.81 cm), 18-gauge needle is used. The point of needle insertion is about 2 inches (5.0 cm) vertical from the point of the keel bone. The needle is held at a 90° angle to the plane of the keel bone. Proper bird positioning is essential for consistent results with this method. As in the anterior approach, incorrect needle insertion may require the bird to be humanely euthanized before fatal hemorrhage occurs.

PROPER BLOOD SAMPLE HANDLING ————-

Once a blood sample has been collected into a syringe, the sample should be carefully transferred to a tube to promote clot formation. Clotting occurs when all the cells in the blood are drawn together by the coagulation process and separates from the fluid portion of the blood (serum).
• The needle should be removed from the syringe before the blood is pushed into the clotting tube (Figure 13). Forcing the blood back through the needle will rupture red blood cells (hemolysis), resulting in a poor quality sample.
• Slowly inject the blood into the clot tube, allowing it to run down the side of the tube, which encourages clot formation. Blood must be placed in the clot tube before the coagulation process begins.
• Do not disturb the blood tubes while the clotting process is occurring. Tubes should be allowed to stay positioned nearly flat (horizontal) to maximize surface area of the clot as it forms . The amount of serum yielded from clotted blood depends on the surface area of the clot. Tubes held upright in the vertical position have little surface area and produce only a small quantity of serum. Use a test tube holder for keeping tubes in this flat position. If a test tube holder is not available, then a block of wood drilled with appropriate sized holes or a wire rack can be used.
• The time required for a clot to form depends on the ambient temperature where samples are kept. The ideal temperature for clot formation is 80 to 100°F (27 to 38°C). At this temperature, serum separation will take approximately 12 to 18 hours for completion. At cooler temperatures, the clotting process is slower and the serum yield is reduced. Blood samples can be damaged and are subject to bacterial contamination if exposed to higher temperatures for longer periods of time. This can occur when blood samples are left in a hot car or in direct sunlight.
Bacteria or mold contamination will cause serum to appear slimy with solid cheese-like particles. Opportunistic microorganisms feed on the antibodies in the serum and lower the amount of antibody measured by the laboratory. If birds are dehydrated (in especially hot weather or due to stress), they produce poor serum samples that are gelled. Additionally, serum from birds after a recent meal appear cloudy due to excess fat in the serum. Lipemic (fatty) samples are not ideal to run in the laboratory, as the fat will interfere with any optical based test or antibody fixing test such as ELISA. Blood in the process of forming a clot should not be frozen. The samples should not be shaken or allowed to roll around. Roughly handled blood samples will yield serum containing the pigments of ruptured red blood cells. This process is called hemolysis and makes serum appear red or pink in color. Hemolysis interferes with laboratory tests measuring antibody levels. Samples containing blood clots should not be mailed, as significant hemolysis can occur en route to the laboratory.

SERUM COLLECTION AND HANDLING ————-

After serum has separated from clotted blood, pour serum out of the clot tube into another collection tube, or tease the clot out of the tube with a wooden stick (such as a toothpick), leaving only serum in the tube. A clot must be handled gently during the process of separating the serum. A good quality serum sample will appear clear to pale yellow in color. Cloudy, slimy, or hemolyzed samples should not be sent to the laboratory.

Sending Serum Samples to the Laboratory ———–

Once serum has successfully separated from a clot, it should be kept cool (45°F or 7°C) and sent immediately to the laboratory. Serum should not be frozen if intended for use within 3 to 5 days. The tubes containing individual bird serum samples should be tightly capped, organized by flock into sealed plastic bags and clearly identified with labels or indelible ink. Styrofoam insulated containers with at least one cold pack should be used for mailing. It is best to avoid mailing serum samples to the laboratory on Thursdays or Fridays, as they will arrive at the laboratory over the weekend. Serum that must be stored for longer periods of time should be frozen at +14°F to -40°F (-10°C to -40°C).
NB-Do NOT send serum samples to the lab that:
• Contain less than 0.25 mL of serum
• Are excessively hemolyzed (red)
• Are excessively lipemic (fatty)
• Contain clots
• Are gelled, slimy or contain cheese-like particles

HISTOPATHOLOGY

Histology refers to the evaluation of cells and tissues using a microscope. As a follow-up to the post-mortem exam, histology can be a valuable tool in assessing flock health. Some poultry diseases can only be diagnosed by histopathology. For example, the clinical presentation of infectious laryngotracheitis virus or wet pox within a flock can be virtually identical, but the diseases cause distinctly different and characteristic histopathologic changes that allow a definitive diagnosis. Successful use of histopathology as a diagnostic practice requires the availability of appropriately selected and preserved samples.

Sample Collection ————

Collect specimens for histopathology as soon as possible after death to avoid deterioration of tissues. Fresh tissue samples from birds humanely euthanized immediately prior to postmortem examination provide the best quality slides. If mortality must be used for tissue collection, they should be determined to be fresh as possible, and not decomposed. Do not collect samples from birds that have been previously frozen. The freeze and thaw processes can disrupt cellular features, leading to poor quality slides. Samples should be collected using a scalpel or razor blade that is sharp and sterile (Figure 5). Avoid using scissors, as they can crush tissue and destroy microscopic details. An individual sample should be no larger than 1 cm3 (1x1x1 cm) to allow for adequate penetration of the tissue with fixative. Larger pieces of tissue will decompose in the center before adequate penetration by the fixative (formalin).

Sample Selection ————

Samples for histopathology should be collected at the time of post-mortem analysis. The selection of samples depends on observations made during the examination. Tumors and other masses, focal discolorations, and organs that are enlarged, atrophied, or otherwise abnormal should be sampled. When a particular disease is suspected based on flock history, tissues associated with that disease may be collected, even if they appear normal (see Table 1). A crosssection of all parts of the affected organ being sampled should be harvested whenever possible. Tissue cut from the margin of the lesion, collecting both affected and normal tissues, is preferred. Whenever possible, collect healthy-appearing tissue of the same for comparison.

Sampling for Specific Diseases

When there is concern for a particular disease based on regional risk, a suspicious result on surveillance testing, or clinical signs in the flock, specific tissues should be collected. Table 1 provides examples of some diseases of concern and the special samples that should be taken.
Disease of Concern ——————–Samples Needed
Gumboro (IBD) ———–• Bursa of Fabricius, Thymus
Infectious Laryngotracheitis ———-• Trachea • Larynx • Conjunctiva
Marek’s Disease———— • Sciatic Nerve • Brain • Eye • Tumors
Wet pox ———-• Trachea • Larynx
Enteritis (coccidia, focal duodenal necrosis)————- • Portions of the gastrointestinal tract affected

Sample Preservation————–

Samples should be promptly submerged in a solution of 10% neutral buffered formalin for preservation. The volume of formalin solution in a single container should be at least 10 times the volume of all tissues. Samples must be fully immersed in the solution to be adequately saturated by fixative to prevent deterioration. Lung tissue and other air-containing tissues may be wrapped gently in absorbent cotton to aide immersion. Gently open the lumen of trachea and intestine samples to release trapped air. After 48 hours in formalin, the tissues are adequately fixed. If necessary for shipping, the formalin can be decanted at this point. Decanted samples should be shipped immediately to minimize the risk of damage to the sample from drying. If samples may be subject to sub-freezing temperatures during shipping, already fixed samples can be decanted, and re-submerged in an “alcoholic formalin.” This will protect against freeze-thaw damage to tissues. For a simple alcoholic formalin mix, combine and pre-mix 6.5 parts pure ethyl alcohol, 2.5 parts distilled water, and 1 part 37% formalin. The formalin-fixed samples can be kept in sealable plastic bags (e.g. Whirl-Pak® bags), or remain in a securely sealed jar with formalin. If the samples are to be mailed to the laboratory, double-bag the sample to prevent leakage. Remember that formalin is a poison and exposure to the liquid or vapor is harmful to humans.

Sample Submission—————

When submitting samples to a diagnostic laboratory, it is important to provide thorough and relevant flock information on the laboratory submission form. Critical information that should accompany all diagnostic sample submissions includes:

• Flock identification and location
• Age of flock
• Date of sample collection
• Tissue(s) collected
• Vaccination program
• Flock history, including description of any clinical signs, production problems, and the present level of mortality
• Special shipping regulations may apply for formalin filled containers
• Locally appropriate biohazard labelling on all transport containers
• Appropriate permits for international shipping (e.g. USDA-APHIS permits) if appropriate This information is vital to the flock veterinarian and diagnostician to make a meaningful interpretation of diagnostic results and provide recommendations to improve flock health and/or production.

Sample Processing

After arrival at the diagnostic laboratory, the formalin preserved tissues are embedded into a paraffin block, then sectioned with a microtome into thin slices. Tissue slices of this thickness (4 micron) are thin enough to be examined by the pathologist under a light microscope. These slices are fixed on glass slides and stained. Various stains can be used to highlight different cell types, or other aspects of the tissue. The most frequently used stain for disease diagnosis is hematoxylin and eosin (H&E) stain

Preparation of Specimen for Disptach

An initial incursion of an exotic disease in most cases, only be confirmed in a reference laboratory through the isolation and identification of the etiologic agent. Thus specimens to be submitted for agent identification should be collected strict aseptically and completely. The following general suggestions are presented as a guide for preparing diagnostic specimens for submission to a diagnostic laboratory.
1. Obtain and record a complete herd history. Information should be submitted on proper forms, when possible the following information should be included.
– Name and address of the owner. – Name, address and phone number of submitter. – A description of animal : Breed, Sex, Peculiarities etc. – Suspected disease or examination requested or both. – Number of animals showing similar symptoms and age of the animals. – Number of animals died due to the condition. – Vaccines administered to the animal(s) from which specimens were collected especially important when examinations for antibodies required. – Date of the first losses and subsequent losses. – The disease symptoms and their duration. – Ration fed. – The general condition of the animal. – A description of the spread of the infection, if in a flock or herd. A diagram of the area is often useful. – Treatment given if any. – Type of housing. – Accessory information like the type of preseravative used for specimens. – An epidemiological assessment, including recent movements into and out of the flock or herd. – Any exposure of the affected poultry or livestock to persons.
2. Be objective and approach the investigation without a preconceived diagnosis.
3. Be alert regarding safety hazards in handling livestock and consider zoonotic potentials e.g.possibilities of rabies as a differential diagnosis should be considered where appropriate.
4. Ensure that prelabeled specimen containers and tubes are available for collection and are scrupulously clean and sterile. The label must include proper identification of the animal and type of specimen.
5. Examination and collection of specimens from live animals or poultry in various stages of clinical disease. Serum, vesicular fluid or tissue, or both, swabs of exudates or lesions or both can be secured from live animals. Serum from apparently healthy exposed animals or poultry can also be helpful. Animals sampled should be permanently identified because it is possible that convalescent serum or sample will be taken in future for comparative purpose.
6. Blood smears should be prepared on clean glass slides. A thin blood film should be made, rapidly dried, and fixed in absolute methanol for 5 minutes. Slides having frosted end should be used and should be identified using a lead pencil .

Collection of Specimens at Necropsy———-

1. Collect specimens from the animal which have undergone minimal putrefaction. 2. Be aware of any safety or biological hazards that necropsy might impose on you and the owner. Availability of a proper and safe disposal site should be considered before begining necropsy. 3. Wear rubber boots, gloves, overalls etc. that can be disinfected or that are disposable. A mask and goggles may be used at the discretion of the diagnostician. 4. Prelabeled specimen containers will help ensure that recommended specimens will be collected. – Use a label that can not be easily destroyed. – Writing should be with pencil or ink that will not smudge or blur when wet. – Use plastic screw-cap container istead of glass container where practical. – Tape the lids of containers tightly so there will be no leakage as well as entry of other material. – Use disposable equipment such as cardboard trays, disposable syringes etc. 5. Have a systematic plan for the necropsy and know what specimens are to be collected before starting the procedure. Be certain to include all lesions for laboratory examination. Body fluids and contents of cysts, abscess, or skin lesions can be collected using a sterile swab. If an animal is presented for euthanasia, collected blood sample before euthanasia. If the animal or bird is presented dead, collect blood from the heart. Make blood smears as per standard method. Ectoparasites should be noted and collected, if present. The collection of specimens based on species rather than a specific disease will be most useful in providing a diagnosis in some diseases. Samples of all lesions should be collected for histological examination. 6. Fluid from any enlarged joints, if found should be aspirated aseptically. 7. Any excess body cavity fluids should be collected aseptically via a syringe. Some other considerations in Specimen collection 1. Two sets of tissues are to be collected. – Fresh tissue for microbiological examination. – Preserved tissue for histological examination. The recommended preservative is 10% neutral buffered formalin. All tissues can be placed in one container, but allow not more than 1 volume of tissue to 10 volumes of formalin. Tissues from organs should be cut perpendicularly to the surface to expose their anatomic structure. The specimen should include affected and surrounding normal tissue. To provide adequate fixation, tissue except brain, should be sliced no more than 3-6 mm thick. Any lymphnodes collected should be incised. Specimens should not be folded or bent the container in which they are fixed. Only wide mouthed containers should be used in this procedure. 2. The initial piece of each organ or lesion should be collected aseptically for microbiology. Tissues for formalin fixation can be collected during the necropsy. 3. Swabs should be sent in appropriate transport medium. The laboratory can assist in the procedure for obtaining the media. 4. Materials submitted for possible virus isolation should be obtained from animals that died and have minimal putrefaction and from animals in the early, acute, febrile phase of illness. Specimens shipped for virology and bacteriology should be shipped refrigerated. If at all possible, the use of dry ice should be avoided because of CO2 will produce acid conditions that will inactivate many viruses. If there is no way to submit the specimens to the laboratory within 48 hours, dry ice must be used. In this case, the specimens must be completely sealed so that there is no contact of the gas emitted by the dry ice with the specimens.

Post necropsy considerations ————

1. Clean and decontaminate all instruments and premises. 2. Record necropsy findings. 3. Dispose of carcass and body parts so as to avoid exposure to other animals and contamination of environment.

Considerations for shipping diagnostic specimens———-
Regulations require that diagnostic specimens transported in interstate or inter country traffic must be packaged and labeled properly. Improper packaging and labeling of diagnostic specimens and other hazardous material can result in unnecessary exposure to postal, shipping, laboratory personnel etc. 1. The specimens must be in securely closed, waterproof primary enclosure such as screw cap container or sealed vial. Be certain that exterior surfaces of the primary containers are decontaminated before shipment/air parcel. 2. Each primary container should be wrapped in sufficient dry absorbent cotton or paper towels to absorb the material in case of breakage. Ideally, the wrapped container should be placed in sealed plastic bags. 3. Pint, quart or half gallon sized paint cans should be used as secondary containers. These cans should have friction type lids and be water proof when hammered closed. The primary container should be padded with more cotton or paper to prevent jarring. A tertiary container, such as large sized version of the secondary cotainer, should be considered if a zoonotic or highly infectious disease is suspected. 4. The sealed secondary of tertiary container should be placed in a shipping container and again packed with material such as paper. The shipping container should be an insulated box with a lid that can be taped shut. A corrugated shipping box, affixed with the proper labels and shipper’s certification, is the final enclosure and contains all other containers. 5. If specimens can be in transit for less than 48 hours, ice packs may be used for cold storage. Frozen “foam ice”, “blue ice” picnic packs, or water frozen in sealed containers may be used. Wet ice, even when wrapped in plastic bags, should be avoided to eliminate the possibility of leakage. 6. Dry ice is the only suitable refrigerant to keep specimens frozen. Shippers must be aware of dry ice restrictions imposed by certain airlines and plan according. 7. Regular mail or airmail shipment should not be used when a exotic infectious animal disease is suspected. Courier service is the appropriate method of shipment. If FMD is considered as a possible diagnosis, a responsible individual should hand carry the specimen to the reference laboratory. 8. If it is not desirable to have the submission form, with the history and other information, within the container. It is preferable to enclose the submission form between the shipping container and the cover of the outside corrugated box. 9. The shipper is responsible for notifying the intended recipient of all information related to transportation arrangements in order to expedite package pickup and delivery to the laboratory. 10. Care must be taken to ensure that a exotic infectious animal disease suspicious package is only opened within the confines of a biosecure facility

Please follow and like us: