Each veterinary diagnostic laboratory offers a unique set of diagnostic tests that is subject to frequent changes as better tests become available. The protocols for sample collection and submission are therefore also subject to change. The practitioner and diagnostic laboratory staff must maintain good communication to complete their diagnostic efforts efficiently and provide optimal service to the client. Practitioners must be specific and clear in their test requests. The laboratory staff can provide guidance when there are questions regarding sample collection and handling, as well as offering assistance in interpretation of test results. Most diagnostic laboratories publish user guidelines with preferred protocols for sample collection and submission, but the following broad recommendations are fairly standard.
Regardless of the type of submission, a detailed case history should be included with the samples to assist laboratory personnel in determining a diagnosis.
A detailed case history should include:
gross appearance (including size and location) of the lesion(s)
previous treatment (if any)
time of recurrence from any previous treatment
morbidity/mortality in the group
If a zoonotic disease is suspected, this should also be clearly indicated on the submission form to alert laboratory personnel. The submission form should be placed in a waterproof bag to protect it from any fluids that might be present in the packaged materials.
Packaging of Samples for Shipment
The shipment of biologic specimens should comply with protocols established by the courier or shipping service used. In some instances, air transport requires compliance with IATA (International Aviation and Transportation Association) regulations for hazardous materials. These regulations include restrictions on the volume of formalin that can be shipped in a container (<1 L in total and <30 mL per jar) and that shipments with fresh tissue samples are clearly labeled as Biological Substance Category B. If high risk, reportable diseases are suspected, it is essential to contact state and/or federal veterinary agencies regarding necessary shipment precautions. Further details can be found at the International Air Transport Association website and the US Postal Service Postal Explorer web page as well as commercial courier websites.
A fundamental approach is to devise a 3-layer barrier to protect the sample. The sample is placed in an appropriate primary container (sealed jar/bag/tube). This is then enclosed in a secondary container, which includes some adsorbent material. Note that items such as syringes, obstetrical gloves, and containers without sealable orifices are not suitable for shipment. Liquid samples should not ship in plastic bags; a sealable jar should be used. Waterproof markers should be used when labeling specimen bags and containers: the contents and patient identification are critical information.
The secondary container is then placed in the shipping box (tertiary container), which often houses coolant packages as well as various cushioning materials (eg, polystyrene foam) to protect the sample. The coolant materials should be sealed in plastic bags to prevent condensation damage. Coolant packs should not be placed directly onto samples, such as tubes of whole blood, that could suffer adverse effects if frozen in transit. Be sure to include the suitably protected submission form. The tertiary container is ideally a sturdy polystyrene refrigerator box or a cardboard box lined with a fitted polystyrene lining. If dry ice is used, this should be noted on the cardboard box label, and the lid should not be sealed with tape. Otherwise, CO2 released from the dry ice could increase pressure and damage the package or contents.
Tissues for microscopic examination collected either via biopsy or during necropsy can be critical to obtaining a diagnosis. Use of this relatively rapid and inexpensive diagnostic technique can often result in substantial savings in time, money, and animal life. The increasing number of immunohistochemical (IHC) tests that can be applied to formalin-fixed tissue has further reinforced the utility of this diagnostic technique.
Autolyzed tissues are generally useless for histopathologic examination; prompt necropsy examination and organ sampling are critical. Tissue should not be frozen before fixation. Other than CNS tissues (see below), samples collected for histology should never be >1 cm thick (preferably 5–7 mm) and must be placed immediately into ≥10 times their volume of phosphate-buffered 10% formalin to ensure adequate fixation.
Tissues collected for histologic examination should be representative of any lesions present and, in the case of cutaneous punch biopsies and biopsies obtained via endoscopic collection, be centered directly on the grossly visible lesions. Wedge biopsies or tissue samples collected at necropsy should include some of the apparently normal surrounding tissue; the interface between normal and abnormal may provide key information. Excisional biopsies of small tumors (<1.5 cm) may be cut in half. Larger tumors may be sliced like bread so that formalin can penetrate to the face of each slice. Alternatively, several representative samples (7 mm wide, including the interface of normal and abnormal) may be collected.
Rather than shipping large volumes of formalin, tissue samples can be fixed in-house for at least 24 hours in an appropriate volume and then transferred into a smaller volume of fresh formalin for shipment. Prolonged fixation can adversely affect immunohistochemistry testing, so samples should be shipped promptly if immunohistochemistry tests are anticipated. Histologic samples should be shipped in unbreakable containers and packed in a manner that prevents spillage or freezing during shipment.
Specific tissues collected at necropsy require additional attention. Because the GI mucosa decomposes rapidly, short sections of gut collected at necropsy should be opened lengthwise to allow adequate fixation. If spinal cord is to be submitted, the dura mater should be carefully incised lengthwise to permit more rapid penetration to the cord.
Fixing the brain poses a special dilemma, especially if a neuroanatomic location of the lesion(s) within the organ could not be determined antemortem. Ideally, a whole, intact, fixed brain is required for complete histopathologic analysis. Immersion of the brain for many days in a very large volume of formalin is required to adequately fix such a specimen, so brains are commonly transported in an only partially fixed state. If the specimen can be shipped by overnight delivery, it may be acceptable to send a chilled, carefully packaged, unfixed brain, which can then be processed at the diagnostic laboratory. Often, the brain is halved longitudinally and one-half sent unfixed (fresh), properly refrigerated, for microbiologic tests, while the other half partially fixes in transit. This method can prove unsatisfactory if a solitary unilateral lesion is involved. Slicing the brain into widths suitable for rapid fixation introduces considerable fixation artifact and should be avoided if possible; fixing the intact/halved brain in a large volume of formalin for >24 hours is preferred.
Any specific agents of interest in the diagnostic investigation should be mentioned on the submission form; some agents have requirements (eg, anaerobic culture, special media) that would not be used in most laboratories unless the pathogen was cited as a differential diagnosis. Laboratory techniques and capabilities for microbiologic examination vary, but most tests rely on either the growth/visualization of intact viable organisms or the detection of the nucleic acids and proteins of these pathogens. Therefore, unfixed specimens (tissue, fluid, etc) should be collected aseptically and shipped promptly to avoid degradation.
If PCR testing is to be performed, it is particularly important to avoid cross-contamination between multiple animals in a submission; this applies to tissues, fluids, and even dissection instruments. Furthermore, swabs destined for PCR analysis should not be placed in agar or charcoal-based transport media. Calcium alginate swabs should be avoided. Instead, cotton or dacron swabs should be shipped in a tube with a few drops of sterile saline or viral transport media.
Some test protocols may permit pooling of organ specimens from an individual, but for the vast majority, it is preferable that each tissue be collected into separate, sterile, clearly labeled bags or tubes for shipping. Gut samples must never be pooled in a container with other tissue samples. Tissues and fluids for most microbiologic assays may be frozen before shipment, but freezing is generally undesirable if samples can be chilled and delivered directly to the laboratory within 24 hours of collection. Exceptions to this rule include analysis for certain toxins, such as those of Clostridium perfringens and C botulinum, in which degradation of the toxin must be prevented by prompt freezing after collection. Adequate coolant should be provided so that samples remain chilled (or frozen) until they reach the laboratory.
Fecal samples for parasitology testing should be submitted chilled in appropriately sealed containers. Freezing may have little impact on routine flotation or sedimentation tests but will negate the possibility of Baermann analysis for nematode larvae. Ectoparasites or nematodes being submitted for identification should be submitted in vials containing 70% alcohol.
If a known toxin is suspected, a specific analysis should always be requested—laboratories cannot just “check for poisoning.” A complete description of clinical and epidemiologic findings may help differentiate poisoning from infectious diseases that can simulate poisoning.
The most critical samples to be collected are generally:
and some exceptions, such as cerebral tissue for cholinesterase analysis
For some investigations, the diagnosis requires analysis of feed or water. If there is doubt about sample submission procedures, the laboratory should be contacted.
Freezing is critical to prevent the degradation of only a few analytes, such as cholinesterase, zinc phosphide, and sodium fluoroacetate (Compound 1080). For most analytes, overnight shipping of chilled specimens will suffice.
The containers for packing and transporting specimens should be free of chemicals. Plastic containers, both bags and jars, are ideal: jars with metal tops should be avoided. Samples should be packed individually and all containers labeled.
If legal action is a possibility, all containers for shipment should be either sealed so that tampering can be detected or hand-carried to the laboratory and a receipt obtained. The chain of custody must be accurately documented.
If feed or water is suspected as the source of poisoning, chilled samples of these and any descriptive feed tag should accompany the tissue samples. If at all possible, a representative composite sample of the feed should be submitted from the suspect lot or shipment (ie, aliquots from the top, middle, and bottom of the feed container).
Routine studies require anticoagulated whole blood and several blood smears. Blood smears should be prepared immediately after the sample has been collected to minimize cell deterioration. Anticoagulated blood should be kept refrigerated; blood smears should not. EDTA is the anticoagulant of choice for a CBC. Blood for coagulation testing should be collected into a blue-top tube, which contains sodium citrate. After mixing, the sample should be centrifuged for 5 minutes, and then plasma should be removed and transferred to a clean tube without anticoagulant. The plasma should be kept frozen until the time of analysis. Whole blood should not be frozen because this causes cell lysis and gross hemolysis, which interfere with testing.
Most clinical chemistry tests require serum, but an occasional test may require plasma. Anticoagulants present in plasma may interfere with tests; therefore, serum should always be submitted unless plasma is specifically requested. Because lipemia can interfere with a number of chemistry tests, dogs and cats should be fasted for 12 hours before samples are collected.
For serum samples, the blood should be drawn into a red-top tube or a separator tube. The sample should be held at room temperature for 20–30 minutes to allow complete clot formation and retraction. Incomplete clot formation may cause the serum to gel due to latent fibrin formation. The clot should be separated from the glass by gently running an applicator stick around the tube walls (“rimming”). The sample should then be centrifuged at high speed (~1,000 g; 2,200 rpm) for 10 minutes. Rough handling of the sample or incomplete separation of erythrocytes from serum may promote hemolysis, which can interfere with certain tests.
If the sample has been collected into a serum separator tube, centrifugation will cause a layer of silicon gel to lodge between the packed cells and the serum. The gel layer should be inspected to ensure the integrity of the barrier, and re-centrifugation is recommended if there is a visible crack in this layer. If a red-top tube has been used, the serum should be removed and transferred to a clean tube to minimize artifacts such as decreased glucose levels. Serum should be refrigerated or frozen until analyzed. Delays in analysis may adversely affect the results for a number of analytes.
Serology generally requires serum, but plasma is often satisfactory. Samples should be collected as described for clinical chemistry tests and should always be free of hemolysis. In some instances, paired samples may be required for an adequate diagnosis. The acute sample should be collected early in the course of the disease and frozen. The convalescent sample should be collected 10–14 days later, and both samples should be forwarded to the laboratory at the same time.
Air-dried smears are usually acceptable. Rapid air drying of smears minimizes cell distortion, thereby enhancing diagnostic quality. However, depending on the method of staining used, some laboratories prefer alcohol-fixed smears. Samples can be obtained by fine-needle aspiration or by scraping. Imprints (touch preparations) of external lesions can also be used, although these tend to have a greater degree of contamination. Aspirated material should always be smeared before air drying. Smears of fluid can be prepared using a traditional blood smearing technique. Highly cellular fluids may be smeared directly; fluids of low cellularity should be centrifuged to concentrate the cells. Thick material or viscous fluid is more readily smeared using a squash technique in which a second glass slide is placed over the aspirated material and then slid rapidly and smoothly down the length of the lower slide.
Blood or cytologic smears should never be mailed to the laboratory in the same package with formalin-fixed tissues because formalin vapors will produce artifacts in the specimen. Many laboratories now offer immunocytochemical testing, and proper handling of cytologic submissions is required for reliable results. Usually air-dried, unfixed smears will suffice, but in some instances, shipping of samples in tubes containing a transport media is recommended.
Analysis of various effusions and fluids usually includes determination of protein content, total cell count, and cytologic examination. Other tests may be performed depending on the source or appearance (eg, chylous fluid) of the effusion. A sample of effusion/fluid should be collected into an EDTA (purple-top) tube for routine analysis. A second sample should be collected into a serum (red-top) tube if any biochemical analyses (eg, triglyceride, cholesterol, lipase for chylous effusions) are to be performed or if a bacterial culture is desired (eg, joint fluid). These samples should be shipped chilled but not frozen.
Smears for cytologic examination (see above) should be prepared from a drop of the fluid immediately after the sample has been collected to minimize cell deterioration and other in vitro artifacts. Samples of CSF should be collected into small EDTA tubes and shipped immediately with high priority; the cytologic value of CSF samples degrades rapidly, and the low cellularity makes examination of direct smears unrewarding. If sufficient CSF is available, then a red-top tube sample may be useful for serology or culture attempts.
Urine is another fluid that suffers rapid degradation. Generally, if urine cannot reach the laboratory within 12 hours of collection, there is likely to be some degree of inaccuracy in results obtained. Thus, these samples must also be shipped chilled and with high priority. Some laboratories accept urine samples with addition of a preservative such as boric acid, but clinicians should contact these labs for specific protocols. (See also Overview of the Urinary System Overview of the Urinary System Primary functions of the urinary system include: 1) excretion of waste products of metabolism; 2) maintenance of a constant extracellular environment through conservation and excretion of water... read more .)
Tests based on the detection of specific genetic features range from karyotype analysis to the identification of specific genes. The laboratory offering the test should be contacted to determine the specifics of sample collection and handling; required samples range from hair to skin or blood. Many blood-based analyses require collection into yellow-topped acid-citrate-dextrose tubes and overnight shipment of the chilled tubes to the laboratory. Tissue samples for genetic analysis should be unfixed and shipped immediately after collection. As with most molecular techniques, aseptic collection and the prevention of cross-contamination between samples is critical for reliable test results.
Communication between the clinician and the veterinary diagnostic laboratory is critical to assure that the appropriate test and sample are used and that results are interpreted correctly.
Samples should be clearly labeled with waterproof marker on appropriate primary containers, then shipped to the laboratory using the "three-layer barrier" format, including a suitably protected submission form detailing case features and specific test requests.