Summary
Good lab results start with maintaining a strong protocol that seeks to minimize pre-analytical effects. This starts with sample collection, understanding the value of different sample types, and handling of samples all the way to the laboratory. Ideal procedures may vary by species and will certainly vary with different reference laboratories, so it is important to establish a standard operating procedure within the clinic and update it regularly. Test procedures performed in-house should also be routinely reviewed. Understanding the basics of reference interval guidelines will enhance your interpretation of test results.
Introduction
Standard procedures to ensure proper sample collection and handling are critical to minimizing preanalytical variables.1-3 Obtaining the best diagnostic results begins with proper sample collection. The choice of the wrong tube or swab can alter the reliability of the diagnostic results. Even if the proper tube is selected, under filling a tube resulting in excess anti-coagulant can also induce alterations. After collection, the samples should be handled with care to minimize additional changes. These procedures include prompt centrifugation, separation of plasma, and proper storage and shipping. Then, once results are received, gauging the values vs. fully vetted reference intervals, when available, is best; when not available, there are other options.
Anti-coagulants
Two main anti-coagulants are commonly used with avian blood samples: EDTA and heparin. EDTA tubes are commonly called lavender top tubes (LTT) and most often used for hematologic tests. Under filling of tubes or the use of incorrect blood tubes (i.e., larger vacutainer type) will result in high concentrations of EDTA which will affect the stability of the sample. Most commonly, the red blood cells will shrink, and thus, affect hematocrit and total counts.1,2 If sample volume is not an issue, practitioners should strongly consider submitting EDTA blood for CBC analysis. EDTA will result in a more stable sample.4-7 There have been only a few reports recommending anti-coagulants for a specific species although it is anecdotally known that EDTA should not be used for some passerine and many reptilian species.5 Harr and coworkers reported no significant differences between EDTA and heparin in samples from macaws and pythons at 3 and 24 hours.8 Guzman et al. reported no differences in WBC and RBC counts between heparin and EDTA in samples from Amazon parrots when processing occurred within 2 hours of sample acquisition.9 Notably, however, thrombocyte clumping can increase in heparinized blood.9
Heparin (green top tubes, GTT) is a commonly used anti-coagulant in human clinical pathology as it often results in the least interference with tests.2 It is commonly used in avian medicine for biochemistry determinations as the collection of serum often results in large fibrin clots and latent fibrin formation which can affect test results and also significantly minimize sample volume, thus, reducing the number of tests which may be performed.10,11 Under filling heparinized tubes also can result in some artifacts. Most commonly, WBC are found to be very fragile, thus, making this an undesirable type of sample for CBC which are not completed in-house.4,5 In addition, blood smears made from heparinized blood often have a bluish background.2
Several manufacturers make blood tubes in an array of sizes—many especially for the pediatric market which are ideal for use in avian and exotic species.3 Ideal fill volumes range from 0.1–1.0 ml. Expiration dates should be monitored. In the event that a tube is under filled, it should be noted at the clinic level so that potential artifacts in test results will be anticipated.
Preparation
Plasma should be separated as soon as possible after collection. Centrifugation speed and time should be controlled. Several small inexpensive centrifuges are commercially available and should be present at the clinic level.
If the plasma is not separated before shipment to the laboratory, two main artifacts are likely possible. First, the extended contact of the plasma with red blood cells will alter some analyte levels. Previously, we have demonstrated glucose drops by 35% after 24 hours and 67% after 48 hours.7 In addition, potassium can change significantly over the first 12 hours.7 This short handling time of avian samples is greatly contrasted by that data observed with mammalian samples where separation times from 2–24 hours is acceptable.2 It is important to note that separation by only centrifugation to trap the RBC in the gel barrier is not acceptable. As changes in analytes will still occur, the plasma should be decanted to an inert tube. Second, shipping of whole blood often results in red cell lysis and sample hemolysis. Hemolysis can also occur when the separated plasma is left in a GTT with a gel barrier and the barrier weakens and allows leakage of the cells into the plasma phase. Even mild hemolysis will affect some analytes. Potassium, phosphorus, albumin, AST, LDH, and protein electrophoresis have all been shown to be affected.10,12,13
Separated plasma should be placed in a size appropriate tube with a secure closure. Large tubes often result in sample loss and loose caps can result in leakage and dehydration. A fresh GTT should not be used for storage as this adds excess heparin to the sample.
For hematology submissions, a peripheral blood smear made during or shortly after sample acquisition is key to best diagnostic results. Delays in smear preparation result in increased smudge cells as well as artifactual changes in WBC and RBC morphology. Such artifacts are greatly enhanced by incorrect anti-coagulant:blood ratio also. Smears may be made via the wedge or cover slip methods.3,5
Storage
For short-term storage before shipment, blood and plasma is best stored under refrigeration.14 For delays of 2 or 3 days or longer, freezing should strongly be considered but only after contacting the reference laboratory for specific recommendations. In some cases, even short-term freezing has been demonstrated to cause some changes.10,12,15 However, repeated freeze-thaw of samples should not occur. Blood smears should always be maintained at room temperature.
Shipping
All samples should be shipped via the recommended shipping company of the reference laboratory. This will expedite the handling of your sample and like provide the best overall shipping service to the lab. The express shipper will provide specific guidelines to ensure proper protection of the sample and minimize the chance of breakage. These guidelines should be met and where possible, exceeded. Cold packs can be included in shipments, but great care should be taken to prevent direct contact of the cold pack with either the blood tubes or blood smears. Helpful hacks include placing blood tubes inside hard plastic box shaped slide holders and insulating further with bubble wrap.
Reference Intervals
In recent years, the American Society for Veterinary Clinical Pathology has adopted reference interval guidelines. Much in alignment with that used in humans, the ASVCP guidelines further acknowledge some of the more unique challenges of exotic species including method differences, sex and seasonal differences, and the difficulties in obtaining large numbers of clinically normal animals of particular species.16 In regard to the latter, guidelines include statistical analyses which address the limitation of lower sample sizes. Ideally, n>120 is a goal but realistically n=40 is more often met. There are rules in working with sample sets of less than 40 specimens. Alternatives, while not embraced by the veterinary community, include an indirect method where animal health is unknown.17 There is value with referencing previous bloodwork on a patient. This “index of individuality” denotes that there are many measurands (hematology, chemistry) that may be more relative to the patient rather than a population of the patients of the same species.18 Importantly, with more reference laboratories working with exotic species and more options for point of care testing, clinicians should insist on moving away from “avian ranges” and to species and method specific intervals. With time, sex and age intervals can also be generated.
References
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2. Young DS, Bermes EW. Specimen collection and other preanalytical variables. In: Burtis CA, Ashwood ER, eds. Tietz Fundamentals of Clinical Chemistry. 5th ed. Philadelphia, PA: WB Saunders Company; 2001:30–54.
3. Cray C, Zaias J. Laboratory procedures. Vet Clin North Am Exot Anim Pract. 2004;7:487–518.
4. Fredlin PJ. Destructive effect of heparin on avian erythrocytes. Av Pathol. 1985;14:531.
5. Campbell TW. Avian hematology. In: Campbell TW, ed. Avian Hematology and Cytology. 2nd ed. Ames, IA: Iowa State Press; 1995:3–19.
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11. Harr KE. Clinical chemistry of companion avian species: a review. Vet Clin Pathol. 2002;31:140–151.
12. Cray C, Rodriguez M, Zaias J. Protein electrophoresis of psittacine plasma. Vet Clin Pathol. 2007;36:64–72.
13. Benson KG, Paul-Murphy J, MacWilliams P. Effects of hemolysis on plasma electrolyte and chemistry values in the common green iguana (Iguana iguana). J Zoo Wildl Med. 1999;30:413–415.
14. Rossing RG, Foster DM. The stability of clinical chemistry specimens during refrigerated storage for 24 hours. Am J Clin Pathol. 1980;73:91–95.
15. Thoresen SI, Tverdal A, Havre G, Morberg H. Effects of storage time and freezing temperature on clinical chemical parameters from canine serum and heparinized plasma. Vet Clin Pathol. 1995;24:129–133.
16. Friedrichs KR, Harr KE, Freeman KP, Szladovits B, Walton RM, Barnhart KF, Blanco-Chavez J. ASVCP reference interval guidelines: determination of de novo references intervals in veterinary species and other related topics. Vet Clin Pathol. 2012;41:441–453.
17. Tang F, Messinger S, Cray C. Use of an indirect sampling method to produce reference intervals for hematologic and biochemical analyses in psittaciform species. J Av Med Surg. 2013;27:194–203.
18. Jones MP, Arheart KL, Cray C. Reference intervals, longitudinal analyses, and index of individuality of commonly measured laboratory values in captive bald eagles (Haliaeetus leucocephalus). J Av Med Surg. 2014;28:118–126.