Sample collection


Close attention must be paid to all aspects of sample collection, handling and storage, to ensure the most accurate clinical pathologic results. This section provides general guidelines and recommendations for optimally collecting and submitting blood samples for clinical pathologic testing. More detailed information on specific recommendations for individual tests can be found under related sections (hematology, hemostasis, urinalysis, chemistry, cytology) and under each test result. This information includes the effects of controllable preanalytical variables such as how you collect, handle and store the sample before submission to the laboratory. Note that this page refers to blood samples, but some chemistry tests can also be performed in ocular fluids.

Collection of blood

  • Blood collection tubes

    Blood collection tubes

    Clean venipuncture is essential to minimize artifactual changes in the results.  Blood should flow freely with minimal interruptions during collection. This is particularly important for hemostasis testing.

  • Anticoagulant
    • Hematology: EDTA is preferred. Always submit a freshly made blood smear (preferably unstained) to optimize examination of red blood cell, white blood cell and platelet morphologic features.
    • Hemostasis: Citrate anticoagulant (blue top) is optimal for coagulation assays, whereas EDTA is required for platelet counts. Strict attention must be paid to collection (correct volume, clean venipuncture as detailed more in the related section).
    • Urinalysis: Samples should be collected into sterile containers. Plastic non-anticoagulant tubes should be avoided (contains crystalline material which introduces artifact); glass non anticoagulant tubes (red top) are fine.
    • Chemistry: Either non-anticoagulant (red top) or heparin anticoagulant (green top can be used). Results from the two differ (e.g. total protein and potassium are higher in plasma and serum, respectively), hence the same sample should be used if monitoring a patient sequentially.
    • Cytology: For fluid specimens (e.g. body cavity fluids), EDTA is preferred (preserves morphologic features). As indicated above, freshly made smears (with details provided regarding the type of smear, i.e. unconcentrated or direct or concentrated/centrifuged or sediment) should be provided with the EDTA. A non-anticoagulant tube can be used as well if culture is anticipated or if chemistry tests are warranted (e.g. measurement of bilirubin in a suspected bile duct rupture). If the fluid is hemorrhagic, collection into  a red top will help distinguish between acute hemorrhage (or a splenic tap for abdominal fluid), which will clot, and pre-existing hemorrhage, which will not clot (blood defibrinates rapidly in body cavities).

Handling and storage

Samples should be handled appropriately after collection

  • Separation: For hemostasis and chemistry testing, the plasma or serum should be separated from cells as soon as possible after sample collection. For serum, this involves allowing the sample to clot (for serum) and centrifuging the sample to separate out the blood components. This can take a while in large animals (horses, ruminants), so rimming of the tube (inserting a wooden stick around the edges to separate the clot from the tube) may help facilitate clotting, which can take up to 30 minutes. The serum or plasma should be separated from cells and placed into a new clot (no anticoagulant) tube, which should labeled as “plasma” or “serum”.  Corvac or serum separator tubes can be used (the gel facilitates separation of serum after centrifugation), but if they are old or damaged, the silicon may allow cellular constituents to leak through into the serum after centrifugation or may allow cells to metabolize glucose, therefore it is best if the serum is still removed and placed into a new clot tube.
  • Labeling: All body fluid samples taken from a patient should be correctly labeled with the patient name or identification and the type of specimen (e.g. serum, plasma, synovial fluid, peritoneal fluid).
  • Storage: All fluid samples should be stored at 4°C until submission. In contrast, slides should not be refrigerated (the cells lyse with storage).


  • Label properly: Make sure all tubes and slides are labeled with patient identification and the contents (e.g. blood, urine).
  • Package appropriately: Make sure the samples are protected from breakage. Cardboard slide boxes frequently break during transit, so if you are using them, wrapping them in bubble wrap or other cushioning material will protect them.
  • Add cool packs: Ship fluid samples on cool packs. The samples should not be in direct contact with the cool pack, but wrapped in paper towels. Direct contact of cells with an ice pack will cause freezing of the sample and cell lysis.
  • Avoid formalin: Do not ship formalin containers in the same package as the slides or tubes. Formalin readily leaks out of containers and affects the quality of the samples.
  • Ship ASAP: The quicker the sample gets to the laboratory, the fewer the false changes in results due to storage.
  • Provide a good history: This includes signalment (species, age, breed, sex of patient), history that may be relevant (e.g. travel, access to toxins, current medications), pertinent clinical signs (e.g. epistaxis) and, for cytology, a good description of the aspirated lesions (including imaging findings), e.g. multiple hypoechoic masses in the liver.

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