Overview of Collection and Submission of Laboratory Samples

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, should 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. The tissues should remain in fixative for ≥24 hr; after this initial fixation, the samples may be placed in a smaller volume of fresh formalin for shipment if necessary. Prolonged fixation can adversely affect IHC testing, so samples should be shipped promptly if IHC tests are anticipated. Histologic samples should be shipped in unbreakable containers and packed in a manner that prevents spillage during shipment. Fixed tissues should be protected from freezing.

Specific tissues collected at necropsy require additional attention. Because the GI mucosa decomposes rapidly, short sections of gut collected at necropsy must 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 hr is preferred.

Any specific agents that are 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; available tests include bacteriologic culture, fungal culture, virus isolation, in-situ hybridization, a variety of PCR methods, fluorescent antibody tests, latex agglutination tests, Western blotting, ELISA, and many others. Most tests, including the newer molecular biology techniques, 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 generally freezing is undesirable if samples can be chilled and delivered directly to the laboratory within 24 hr. 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 refrigerant should be provided so that samples remain chilled (or frozen) until they reach the laboratory.

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. For a list of appropriate samples for many of the more common toxicities, see Guidelines for Submitting Samples for Toxicologic Examination. The most critical samples to be collected are generally stomach contents, liver, kidney, whole blood, plasma/serum, and urine, but exceptions exist, 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.

Tissues or fluids for chemical analysis should be as fresh as possible and kept refrigerated. For some analyses, freezing is critical to prevent degradation of volatile chemicals, and in rare instances a chemical preservative is required.

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, 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. In some instances, if an adequate amount of involved feed is available, some of it may be fed to experimental animals in an effort to reproduce the signs and lesions seen in the field cases.

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 because it best preserves the cellular components of the blood and prevents platelet aggregation. 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 min, 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 hr 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 min 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 min. 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. Serum should be refrigerated or frozen until analyzed.

Many commercial laboratories provide sample containers and mailers.

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.

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.