College of American Pathologists
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  Q & A





September 2009

Fredrick L. Kiechle, MD, PhD

Question Q. We recently had several physicians question our high-density lipoprotein values, which changed from one testing date to another. For example, on March 7, 2008 a patient had an HDL of 26 mg/dL. The patient was retested June 6, 2008 and his HDL was 33 mg/dL. The affiliated hospital repeated several tests, and we correlated very well. These physicians believe that since HDL is genetic, the results should not vary that much. I have looked in the Tietz Textbook of Clinical Chemistry, and it appears there can be wide variation. Can you clarify this so I can give our physicians the correct information?

A. There are multiple sources of variation that could contribute to the apparent change in the two HDL-C assay results. First, the total error goal for HDL-C in the usual range of HDL-C (>42 mg/dL) is 13 percent, which most current HDL-C assays now achieve. Total error is the sum of any error from any analytical bias, which is recommended to be less than 10 percent, plus imprecision, which in general is recommended to have a coefficient of variation of less than four percent. A slightly less stringent fixed criteria of standard deviation less than 1.7 mg/dL is used for the allowable imprecision for those values of HDL-C less than 42 mg/dL. If we assume the first assay result was accurate, the 95 percent confidence interval for the reanalysis would be between 20.1 mg/dL and 31.9 mg/dL. Thus, the difference between the two assay results just exceeds the total error goal for HDL-C.

Though HDL-C levels are in large part genetically determined, the average biological variation for HDL-C is approximately 7.5 percent. However, it can be much larger. Biological variation is one of the main reasons the National Cholesterol Education Program’s guidelines recommend that multiple measurements be done for any lipid and lipoprotein parameters before a treatment approach is selected. Common sources of biological variation that can quickly and significantly alter HDL-C are the presence of acute illness; changes in weight, diet, alcohol use, or smoking habits; or the use of common medications that can inadvertently alter lipids.

Another frequent preanalytical source of variation in lipids and lipoprotein measurement is whether a fasting sample was collected. Though HDL-C does not change as much as the other lipid and lipoprotein parameters with fasting, it can change significantly in some individuals in the immediate post-prandial period. This may be particularly true for the commonly used direct HDL-C assays, which in some circumstances may measure cholesterol on some non-HDL fractions.

Another consideration is that although most HDL-C assays are now fairly well standardized, there can still be considerable analytical variation between the different types of HDL-C assays, particularly for dyslipidemic samples. In a recent CAP Survey on fresh frozen serum, most of the assays yielded results within five percent of each other, but the difference between the highest and lowest result was as large as 15 percent. When interpreting any apparent change in HDL-C and other lipid and lipoprotein test results, it is therefore important to consider also differences in the assay type—there are now more than seven different types of HDL-C assays—or changes in assay generation, or both, as a source of variation.

Finally, it is important to note that both assay results still fall well below the desirable HDL-C range of 40 mg/dL for men and 50 mg/dL for women. Thus, the apparent change in the two results would not affect the calculation of cardiovascular risk assessment in this patient.


  1. Caudill SP, Cooper GR, Smith SJ, et al. Assessment of current National Cholesterol Education Program guidelines for total cholesterol, triglyceride, HDL-cholesterol, and LDL-cholesterol measurements. Clin Chem. 1998;44:1650–1658.

  2. Warnick GR, Wood PD. National Cholesterol Education Program recommendations for measurement of high-density lipoprotein cholesterol: executive summary. The Na-tional Cholesterol Education Program Working Group on Lipoprotein Measurement. Clin Chem. 1995;41:1427–1433.

Alan T. Remaley, MD, PhD
National Institutes of Health
Department of Laboratory Medicine
Bethesda, Md.

Member, CAP Chemistry
Resource Committee

Question Q. If you are performing routine PT/INR or APTT testing, or both, and your result reports out as a critical high value (for example, INR = 8.0 or APTT = 195 seconds), is it acceptable to check the tube for clots by placing a wooden applicator stick into the blood sample and then respinning the tube to sediment the cells and repeating the tests? Some of our staff believe that spinning the tube a second time should not be done because doing so may cause additional trauma to the red cells and release factors that may affect the clotting time.

A. The differential for a critically high PT/INR or APTT result includes congenital or acquired coagulation factor deficiencies, acquired coagulation inhibitors, anticoagulant therapies, preanalytical variables, instrument or operator errors, and, when data are viewed via a downstream information system, postanalytical errors. It is the responsibility of the clinical laboratory staff to investigate for preanalytical and analytical causes of inaccurate critical values before reporting the values to health care providers to prevent providers from taking incorrect actions that may adversely affect patients’ health.

Since most laboratories perform PT/INR and APTT testing in singlet, the first step would be to repeat the critical value test immediately. If the repeat results are markedly different, then the technologist should verify that reagents have not expired, volumes are adequate, and values for two levels of control are within acceptable ranges. If the repeat patient results are consistently abnormal, reagent/instrument problems should still be considered, but in addition the blood sample should be examined for adequate volume and clots and other preanalytical vari-ables should begin to be considered.

Adequate fill volume (≥90 percent of expected) and collection of blood in a blue-top tube containing buffered sodium citrate (.105–.109 µm) should be confirmed when the laboratory receives specimens for coagulation testing. Underfilled tubes increase the plasma citrate concentration, which may lead to incomplete recalcification and prolonged clotting times.1 Plasma from an EDTA tube will also produce markedly pro-longed clotting times because of excess calcium chelation capacity. Obtaining an elevated potassium plasma concentration will confirm this sample collection mistake. Performing PT/INR or APTT on a specimen collected in a tube designed to produce serum will produce critical PT, APTT, thrombin time, and fibrinogen results because of consumption of multiple clotting factors and fibrinogen ex vivo. In addition, to avoid cold activation of factor VII or cryoprecipitation of von Willebrand factor and factor VIII, coagulation tubes should not be stored on ice or refrigerated before testing.1 Prolonged storage at ambient temperature of a capped, adequately filled, citrated tube is another important source of preanalytical variation. Clinical research has validated analytespecific maximum storage times for PT/INR (24 hours), APTT (four hours), and heparin monitoring (one hour).1

Screening for clots by manually decapping the collection tube, stirring with two wooden applicators, and examining the sticks for adherent red thrombi is an accepted technique but one that lacks validation of its sensitivity and specificity to detect unacceptable samples of citrated blood. Eschwege reported another sensitive technique for detecting small, but significant, clots.2 After centrifugation and removal of plasma, the cell pellet was poured through a piece of gauze and examined for macroscopic clots. Eight of 1,334 specimens contained clots (0.6 percent), pro-ducing elevated factor V activity, decreased fibrinogen, and shorter APTT when compared with results obtained on a new sample.

Following centrifugation, plasma should be inspected for gross hemolysis, which can produce shortened APTT and prolonged PT/INR re-sults when red cell lysis is ≥0.9 percent, corresponding to 1.7 g/L.3 Rather than applying a correction factor for hemolysis, the authors recom-mend obtaining a new sample. Hyperlipidemia or hyperbilirubinemia, or both, may also be criteria for rejecting a sample if they potentially produce analytical interference with the clot detection method employed.

In laboratories that use robotic coagulation lines, centrifugation, decapping, or cap-piercing steps are automated, and manual inspection for hemolysis and clots is performed post-analysis.

Before freezing citrated plasma for later batch testing or specialized coagulation testing, or for transport to a reference laboratory, steps must be taken to minimize platelet contamination (platelet count <10 x 109/L) to prevent shortening of subsequent clotbased tests due to platelet membrane fragments.1 Double centrifugation or filtration through a 0.22-µm filter are effective methods of reducing platelet contamination. However, Sheppard, et al., reported absorption of factors V and VIII that was after filtration, leading to spuriously prolonged PT and APTT and positive lupus anticoagulation results. The defective filters were lotspecific. Others have reported filter absorption of selected hemostasis factors,4 and the Clinical and Laboratory Standards Institute discourages filtration to obtain platelet-poor plasma.1

Since I could not find data addressing the effect of stirring the cell pellet followed by centrifugation on PT/INR and APTT results, our laboratory supervisor compared results from five plasma samples with prolonged PT/INR and APTTs obtained before and after checking for clots with wooden applicator sticks. Using a mechanical clot detection method, there was no difference. However, this is not an adequate study to validate the procedure used in the reader’s laboratory, and I would recommend repeating critical value PT/INR and APTT results before checking for clots rather than checking for clots and repeating PT/INR and APTT after a second centrifugation.


  1. Arkin C, et al. Collection, transport, and processing of blood specimens for testing plasma-based coagulation assays and molecular hemostasis assays; approved guideline—5th ed. Wayne, Pa.: Clinical and Laboratory Standards Institute. 2007.

  2. Eschwege V. Overestimation of plasma level of factor V coagulant activity due to unrecognised preanalytical coagulation. Thromb Haemost. 2004;91:827–828.

  3. Lippi G, Montagnana M, Salvagno GL, et al. Interference of blood cell lysis on routine coagulation testing. Arch Pathol Lab Med. 2006;130:181–184.

  4. Favaloro E. Preanalytical variables in coagulation testing. Blood Coagul Fibrinolysis. 2007;18:86–89.

Charles S. Eby, MD
Associate Professor, Division of
Laboratory and Genomic Medicine
Department of Pathology
and Immunology
Washington University
School of Medicine
St. Louis, Mo.

Vice Chair, CAP Coagulation
Resource Committee

Dr. Kiechle is medical director of clinical pathology, Memorial Healthcare, Hollywood, Fla.