BIOPRESERVATION AND BIOBANKING Volume 12, Number 1, 2014 ª Mary Ann Liebert, Inc. DOI: 10.1089/bio.2013.0043
Processing of Diagnostic Blood Specimens: Is It Really Necessary to Mix Primary Blood Tubes after Collection with Evacuated Tube System? Gabriel Lima-Oliveira,1,2 Giuseppe Lippi,3 Gian Luca Salvagno,1 Giorgio Brocco,1 Stefania Gaino,1 Francesco Dima,1 Waldemar Volaski,2 Fabiane Gomes Rego,2 Geraldo Picheth,2 and Gian Cesare Guidi1,2
Background: The preanalytical phase is considered the most vulnerable phase in biopreservation, biobanking, and laboratory diagnostics. Accurate mixing after blood collection is claimed to be important and recommended by the manufacturers. Objective: To evaluate whether it is really necessary to mix the primary blood tubes immediately after blood collection by means of evacuated tube systems. Material and Methods: Blood from 300 outpatients was equally and randomly divided into three groups: G1, sodium citrate vacuum tubes; G2, lithium heparin vacuum tubes; and G3, K2EDTA vacuum tubes. All vacuum tubes were processed using three different procedures. Procedure 1: Gold Standard (P1): All specimens mixed gently and carefully by inverting five times as recommended; Procedure 2: Rest time (P2): All specimens remained 5 min in the upright position, followed by gentle careful mixing by inverting five times; Procedure 3: No mix (P3): All specimens were left in upright position without mixing afterwards. The influence of the primary mixing tube procedure was evaluated for clinical chemistry, hematology, and coagulation parameters by paired t-test. The bias from the mixing procedure was also compared with quality specifications derived from biological variation. Results: Significant differences ( p < 0.017) were found for: i) red blood cell count and hematocrit when P1 was compared with P2; ii) alanine aminotransferase and erythrocyte sedimentation rate when P1 was compared with P3; iii) red blood cell count, hematocrit, and hemolysis index when P2 was compared with P3. Surprisingly, clinically significant differences were found only for sodium when P1 was compared with P2, and P1 was compared with P3. No fibrin filaments or microclots were observed in any samples. Conclusion: Primary blood tubes mixing after collection with evacuated tube system appears to be unnecessary.
key factors in maintaining quality of biospecimens.2 Presently, quality managers focus their attention on the preanalytical phase.3–11 Nevertheless, the manually intensive activities of the preanalytical phase are still considered the most vulnerable steps in biopreservation, biobanking, and laboratory diagnostics. In this field of activity, accurate mixing of tubes with anticoagulant or clot activator additives after blood collection is considered essential and is widely recommended by manufacturers’ datasheets.12,13 Parenmark and Landberg14 recently released a convincing paradigm about the mixing procedure for diagnostic blood specimens. They showed that: i) mixing blood samples immediately after collection may be not mandatory for all types of tubes; and ii) instant mixing may produce spurious
Introduction
B
iomarkers are measurable characteristics represented by proteins, enzymes, ions, and/or macromolecules and cells, which can be used to screen, diagnose, and monitor diseases. It is well known that the process of finding and validating suitable biomarkers is important for clinical research and disease management. Researchers, physicians, and laboratory managers looking for biomarkers may save a large amount of time and money when samples from biobanks are obtained through use of a quality management system.1 Indeed, sample collection and processing are essential steps for biopreservation and biobanking, so that the control of standardized preanalytical conditions are 1
Laboratory of Clinical Biochemistry, Department of Life and Reproduction Sciences, University of Verona, Verona, Italy. Post-Graduate Program of Pharmaceutical Sciences, Department of Medical Pathology Federal University of Parana, Curitiba, Parana, Brazil. 3 Pathology and Laboratory Medicine, Clinical Chemistry and Hematology Laboratory, Academic Hospital of Parma, Parma, Italy. 2
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LIMA-OLIVEIRA ET AL.
hemolysis and thereby introduce a bias for those parameters that are most susceptible to erythrocyte injury.14 The authors also concluded that their study was limited, and results could only be considered as preliminary findings. The main drawback in the outcome of this study was that mixing was performed by putting the tube on a horizontal mixing tray for 1 min (inverting 15 times). This procedure is however different from the manual mixing procedure that is conventionally used by phlebotomists. On this basis alone, the conclusion of increased hemolysis due to immediate mixing could be misleading and cannot be generalized; iii) immediate mixing resulted in a higher plasma hemolysis index in citrate tubes than in heparin tubes, a rather odd finding. Despite this, there was no significant difference between immediate mixing versus 5 min rest before mixing for prothrombin time (PT) and activated partial thromboplastin time (aPTT) results (nor any significant difference in potassium [K] and lactate dehydrogenase [LDH] in heparin tubes). Therefore, this single, though relevant, study provides scant support for an adverse impact of immediate mixing. Relevant to this point, the ISO 15189 standard states that necessary improvements and potential sources of nonconformities, either technical or concerning the quality management system, shall be identified and all laboratory processes shall be validated.15 The aim of the present study was to evaluate whether it is really necessary to mix the primary blood tubes immediately after blood collection using evacuated tube systems.
Europe, Leuven, Belgium) directly into three vacuum tubes, as follows: G1) three 3.6 mL vacuum tubes containing 0.4 mL of buffered sodium citrate (9NC) 0.109 mol/L: 3.2 W/V% (Terumo Europe, Leuven, Belgium); G2) three 3.5 mL vacuum tubes with 52.5 USP U of lithium heparin and gel separator (Terumo Europe); and G3) three 3.0 mL vacuum tubes containing 5.9 mg K2EDTA (Terumo Europe). To eliminate any potential interference due to either the contact phase or the tissue factor, approximately 2 mL of blood was preliminarily collected in a discard tube without additive (Vacuette, Greiner Bio-One GmbH, Kremsmu¨nster, Austria). Blood collection was standardized, including the use of needles and vacuum tubes of the same lot.
Materials and Methods
Procedure 3—No mixing. All diagnostic blood specimens were left in the upright position (i.e., tube stopper at the top), without mixing afterwards. Centrifugation. All samples were centrifuged immediately within 10 min after blood collection as follows: i) sodium citrate vacuum tubes at 1500 g for 15 min at room temperature; and ii) lithium heparin vacuum tubes at 1200 g for 10 min at room temperature. All plasma citrate samples were separated, and because shortening of aPTT in thawed samples may also be caused by residual platelet debris in the thawed sample, a re-centrifugation was done and the sample was then aliquoted. This is the usual practice, to ensure platelet-free citrate samples for freezing. Diagnostic blood specimens storage. All plasma samples, both from citrate and lithium heparin-vacuum tubes were stored in aliquots and kept frozen at - 70C until measurement (22 days after the first blood collection). Samples did not show any signs of hemolysis by visual inspection. No specimen was discarded due to unsatisfactory attempts, difficult venous access, missing veins, manifest hemolysis, or lipemia.
Study design Three hundred adult outpatients of both sexes were invited by the Laboratory of Clinical Biochemistry, (132 males, 168 females, mean age – SD 41 – 3), Department of Life and Reproduction Sciences, University of Verona, Italy, to participate in the study, by allowing three extra tubes to be drawn (*12 mL of additional blood over the normal amount collected by the phlebotomist for routine analysis). These volunteers were equally and randomly divided into three groups (Parenmark protocol)14 as follows: i) G1 sodium citrate vacuum tubes; ii) G2 lithium heparin vacuum tubes; and iii) G3 K2EDTA vacuum tubes. Each volunteer agreed through informed consent to be enrolled in this study, which was also carried out in agreement with the Declaration of Helsinki and under the terms of all relevant local regulations.
Collection of diagnostic blood specimens The collection of blood specimens was performed from 8:00 to 9:00 AM, over a period of 3 weeks by a single expert phlebotomist, according to the recommendations of the Clinical Laboratory Standard Institute (CLSI).16 All volunteers, after 12-hours fasting, were maintained in a sitting position for 15 min prior to phlebotomy in order to eliminate possible interference of blood distribution due to posture.17 After this time frame, a radial vein was located on the right forearm by a subcutaneous tissue transilluminator device (Venosco´pio IV plus, Duan do Brasil, Sao Paulo, Brazil) without tourniquet, to prevent interference from venous stasis,18–20 and approximately 12 mL of blood was collected by venipuncture with a 20-gauge straight needle (Terumo
Processing of diagnostic blood specimens Primary blood tube mixing. All vacuum tubes from G1, G2, and G3 were processed through three different procedures. Procedure 1—Gold Standard.21,22 All diagnostic blood spec-
imens were mixed gently and carefully by inverting five times, —immediately after blood collection—as recommended by the manufacturer.23–25 Procedure 2—Rest time. All diagnostic blood specimens after collection had a 5 min rest in the upright position (i.e., tube stopper at the top), followed by gentle and careful mixing by inverting five times .
Laboratory testing All laboratory tests were performed in duplicate to minimize the analytical coefficient of variation (CVa), so the CVa from analyses in duplicate is smaller than the CVa from internal quality control shown in Table 1. Assessment of hematological parameters. All samples were processed for routine hematological testing immediately after collection ( < 15 min) on the same Sysmex XE2100D, Automated Hematology Analyzer (Sysmex Corporation, Kobe, Japan). The parameters tested included
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GLU* COL** HDL** TG* TP** ALB** CRP* UREA* CRE** UA** ALP* AMYL* AMY-P* AST* ALT** GGT** LDH** CK* BT* BD** P** Ca** Mg* Fe** Na* K** Cl** Lipase* HI**
Tests
(mmol/L) (mmol/L) (mmol/L) (mmol/L) (g/L) (g/L) (mg/L) (mmol/L) (mmol/L) (mmol/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mkat/L) NA
(Units)
2.2 4.0 5.2 10.7 1.2 1.3 21.8 5.5 3.8 4.9 6.4 7.4 8.0 5.4 12.0 10.8 4.3 11.5 11.4 14.2 3.2 0.8 1.8 8.8 0.3 1.8 0.5 10.1 NA
1.5 1.8 4.3 2.0 1.2 1.2 2.2 2.6 2.5 1.0 2.7 0.8 2.2 1.2 1.3 1.6 1.5 3.3 2.2 1.8 3.0 0.7 1.2 2.6 1.0 1.5 1.0 2.8 NA
6.10 [5.33 – 8.71] 4.64 – 0.97 1.38 – 0.30 1.20 [0.85 – 1.60] 73.2 – 5.87 41.5 – 3.37 2.20 [1.40 – 3.70] 6.82 [5.96 – 8.32] 83.1 – 17.7 0.34 – 0.08 1.10 [0.90 – 1.34] 1.02 [0.80 – 1.29] 0.37 [0.34 – 0.48] 0.31 [0.27 – 0.46] 0.31 – 0.09 0.85 – 0.18 5.66 – 0.26 1.41 [1.12 – 1.97] 9.40 [7.70 – 14.7] 3.93 – 2.05 0.98 – 0.18 2.36 – 0.08 0.78 [0.76 – 0.84] 79.6 – 25.9 141 [139 – 142] 4.40 – 0.52 101 – 2.66 0.54 [0.44 – 0.83] 4.3 – 1.8
Procedure 1 Gold Standard
Procedure 3 No mix
6.16 [5.33 – 8.55] 4.64 – 0.95 1.38 – 0.30 1.22 [0.85 – 1.57] 73.2 – 5.72 41.5 – 3.24 2.20 [1.30 – 3.70] 6.68 [6.07 – 8.42] 81.3 – 17.7 0.34 – 0.08 1.10 [0.90 – 1.34] 1.00 [0.78 – 1.29] 0.36 [0.34 – 0.46] 0.31 [0.29 – 0.44] 0.30 – 0.10 0.85 – 0.18 5.61 – 0.26 1.39 [1.12 – 1.97] 9.75 [7.70 – 14.0] 4.10 – 2.05 0.98 – 0.17 2.36 – 0.09 0.78 [0.75 – 0.85] 79.0 – 25.8 141 [139 – 142] 4.39 – 0.53 101 – 2.63 0.54 [0.44 – 0.83] 4.0 – 2.8
Clinical chemistry 6.10 [5.22 – 8.49] 4.64 – 0.94 1.38 – 0.30 1.18 [0.85 – 1.57] 72.8 – 6.13 41.2 – 3.36 2.20 [1.40 – 3.70] 6.78 [6.10 – 8.71] 82.2 – 16.8 0.34 – 0.08 1.10 [0.88 – 1.36] 1.00 [0.78 – 1.28] 0.36 [0.34 – 0.46] 0.31 [0.29 – 0.42] 0.30 – 0.09 0.86 – 0.18 5.64 – 0.26 1.41 [1.16 – 1.96] 9.58 [7.70 – 13.7] 3.93 – 2.05 0.97 – 0.16 2.36 – 0.10 0.77 [0.74 – 0.85] 79.0 – 25.1 140 [139 – 141] 4.39 – 0.52 101 – 2.75 0.54 [0.44 – 0.85] 4.2 – 2.0
Procedure 2 Rest time
vs
vs
vs
Procedure 2 Rest time
0.0 0.0 0.0 1.7 0.6 0.7 0.0 0.6 1.1 0.0 0.0 2.0 2.7 0.0 3.2 - 1.2 0.4 0.0 - 1.9 0.0 1.0 0.0 1.3 0.8 0.7 0.2 0.0 0.0 2.3
mean % difference
0.1255 0.3425 1.000 0.0294 0.0905 0.0863 0.0975 0.2377 0.2189 0.8534 0.2153 0.2759 0.1096 0.3026 0.1036 0.4818 0.8063 0.4246 0.0985 0.3855 0.2001 0.5161 0.9045 0.2048 0.1121 0.6514 0.1429 0.8211 0.6669
P value
1.0 0.0 0.0 1.6 0.0 0.0 0.0 - 2.1 - 2.2 0.0 0.0 - 2.0 - 2.8 0.0 - 3.3 0.0 - 0.9 - 1.4 3.6 4.2 0.0 0.0 0.0 - 0.8 0.0 - 0.2 0.0 0.0 - 7.5
mean % difference
0.7747 0.4731 0.1716 0.4032 0.7746 0.6001 0.9052 0.6468 0.1953 0.4818 0.6675 1.000 0.0708 0.6444 0.0150 0.7899 0.1851 0.7432 0.5987 0.7551 0.5463 0.4909 0.9430 0.0465 0.3363 0.3728 0.1628 0.5877 0.6012
P value
- 1.0 0.0 0.0 - 3.4 - 0.6 - 0.7 0.0 1.5 1.1 0.0 0.0 0.0 0.0 0.0 0.0 1.2 0.5 1.4 - 1.8 - 4.3 - 1.0 0.0 - 1.3 0.0 20.7 0.0 0.0 0.0 4.8
mean % difference
(continued)
0.0498 0.8233 0.1716 0.2350 0.1115 0.1863 0.1589 0.6871 0.5725 0.6669 0.3778 0.2555 0.7768 0.4070 0.1868 0.1036 0.1757 0.9638 0.2005 0.3306 0.0841 0.8097 0.8864 1.0000 0.1161 0.9534 0.6720 0.5297 0.8072
P value
Procedure 2—Rest time Procedure 3—No mix Procedure 3—No mix
Procedure 1 Gold Standard
Procedure 1 Gold Standard
Effect of Three Different Procedures of Primary Blood Vacuum Tubes Mixing in Laboratory Testing
Desirable CVa Bias (%)29 (%)
Table 1.
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1.3 1.2 3.6 7.0 2.0 3.0 NA
4.64 [4.34 – 4.74] 137 [129 – 148] 43.5 – 4.50 96.1 [93.5 – 98.1] 13.1 – 0.63 57.6 – 3.70 6.14 – 1.36 2.96 [2.50 – 3.92] 1.99 – 0.47 0.41 – 0.14 0.20 – 0.03 0.03 – 0.01 0.13 – 0.01 233 – 13.3 9.05 [8.70 – 9.70] 24.0 [17.2 – 34.8]
31.4 – 10.7 38.9 [35.7 – 44.2] 3.08 – 0.15 34.0 [29.5 – 45.0] 46.0 [39.0 – 52.0] 89.0 [82.0 – 102.0] 2.0 [2.0 – 3.0]
31.1 – 10.5 38.6 [36.0 – 45.4] 3.13 – 0.16 34.0 [29.0 – 46.0] 46.0 [38.0 – 55.0] 89.0 [83.0 – 102.0] 3.0 [2.0 – 4.0]
Procedure 2 Rest time
4.67 [4.36 – 4.87] 138 [129 – 150] 43.9 – 1.02 96.2 [93.2 – 98.1] 13.1 – 0.63 58.9 – 3.88 5.97 – 1.36 3.01 [2.50 – 3.87] 1.95 – 0.47 0.41 – 0.15 0.20 – 0.03 0.03 – 0.01 0.13 – 0.01 232 – 13.5 9.10 [8.62 – 9.48] 24.5 [17.2 – 38.0]
Procedure 1 Gold Standard
31.6 – 11.0 38.5 [36.6 – 45.6] 3.10 – 0.16 34.0 [29.0 – 44.5] 46.0 [35.0 – 52.0] 87.0 [81.0 – 102.0] 4.0 [2.0 – 4.0]
Coagulation
4.68 [4.35 – 4.80] 138 [130 – 149] 43.8 – 1.01 96.2 [93.4 – 98.1] 13.1 – 0.63 58.6 – 3.66 6.10 – 1.35 2.99 [2.42 – 3.87] 1.96 – 0.52 0.41 – 0.12 0.21 – 0.03 0.03 – 0.01 0.13 – 0.01 231 – 12.6 9.05 [8.72 – 9.48] 24.0 [18.0 – 35.0]
Hematology
Procedure 3 No mix
vs
vs
vs
Procedure 2 Rest time
- 1.0 - 0.8 1.6 0.0 0.0 0.0 33.3
0.6 0.7 0.9 0.1 0.0 2.2 - 2.8 1.7 - 2.0 0.0 0.0 0.0 0.0 - 0.4 0.6 2.0
mean % difference
0.0790 0.6493 0.2931 0.1195 0.0724 0.7315 0.3360
0.0037 0.2115 0.0012 0.5648 0.8037 0.2835 0.1442 0.9358 0.0545 0.4540 0.7894 0.2674 0.2538 0.6843 0.2756 0.0676
P value
1.6 - 0.3 - 1.0 0.0 0.0 - 2.3 25
0.2 0.0 - 0.2 0.0 0.0 - 0.5 2.1 - 0.7 0.5 0.0 4.8 0.0 0.0 - 0.4 - 0.6 - 2.1
mean % difference
0.0619 0.9224 0.4396 0.0315 0.1006 0.0804 0.1839
0.5374 0.0664 0.4540 0.3194 0.2044 0.8755 0.2107 0.7627 0.6736 0.7524 0.2407 0.0375 0.8098 0.6653 0.1311 0.0033
P value
- 0.6 1.0 - 0.6 0.0 0.0 2.2 - 100
- 0.9 - 0.7 - 0.7 - 0.1 0.0 - 1.7 0.6 - 1.0 1.5 0.0 - 5.0 0.0 0.0 0.9 0.0 0.0
mean % difference
0.4059 0.2872 0.7732 0.6444 0.2548 0.0697 0.0106
0.0068 0.3210 0.0051 0.1232 0.3590 0.3143 0.3869 0.6011 0.2055 0.7250 0.2813 0.3590 0.3665 0.2877 0.5752 0.7272
P value
Procedure 2—Rest time Procedure 3—No mix Procedure 3—No mix
Procedure 1 Gold Standard
Procedure 1 Gold Standard
*Non-normal distribution; the values are presented as median [interquartile range]; P value represents the significance by Wilcoxon ranked-pairs test. **Normal distribution; the values are presented as mean – standard deviation; P value represents the significance by paired Student’s t-test. The bold p values are statistically significant ( p < 0.017),29 according to Bonferroni correction for multiple comparisons. Mean % difference in bold was higher than the desirable bias. CVa (%): Analytical coefficient (within-run precision), by internal quality control on the respective instrument.
2.0 2.3 4.8 15.9 13.9 4.0 NA
(s) (s) (g/L) (%) (%) (%) NA
PT** aPTT* FIB** PS* PC* AT* HI*
1.5 1.0 1.5 1.0 2.0 6.5 3.0 8.0 8.0 9.8 15.5 9.7 7.8 4.0 1.2 8.9
Desirable CVa Bias (%)29 (%)
(10 / L) 1.7 (g/L) 1.8 (%) 1.7 (fL) 1.2 (%) 1.7 (109/ L) 7.8 (109/ L) 5.6 9.1 (106/ L) (106/ L) 7.4 (106/ L) 13.2 (106/ L) 19.8 15.4 (106/ L) (106/ L) NA 5.9 (109/ L) (fL) 2.3 mm/h NA
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(Units)
RBC* HGB* HCT** MCV* RDW** Retic** WBC** NEU* LYMP** MONO** EOS** BASO** LUC** PLT** MPV* ESR*
Tests
Table 1. (Continued)
MIXING OF VACUUM TUBES
red blood cell count (RBC), hemoglobin (HGB), hematocrit (HCT), mean corpuscular volume (MCV), RBC distribution width (RDW), reticulocytes (Retic), white blood cells (WBC) count and differential, including neutrophils (NEU), lymphocytes (LYMP), monocytes (MONO), eosinophils (EOS), and basophils (BASO), platelet count (PLT), and mean platelet volume (MPV). The instrument had been previously calibrated against appropriate proprietary reference standard material and verified with the use of proprietary controls. The erythrocyte sedimentation rate (ESR) was measured in duplicate in the same TEST 1 YDL (Alifax, Padova, Italy) immediately after routine hematological testing on Sysmex XE-2100D. Clinical chemistry tests. All frozen lithium heparin aliquots were thawed at the same time. The routine clinical chemistry tests were performed on the same instrument Cobas 6000 < c501 > module (Roche Diagnostics GmbH, Penzberg, Germany), according to the manufacturer’s specifications and using proprietary reagents. The panel of tests included the following: glucose (GLU), total cholesterol (COL), high-density lipoprotein cholesterol (HDL), triglycerides (TG), total protein (TP), albumin (ALB), Creactive protein (CRP), urea, creatinine (CRE), uric acid (UA), alkaline phosphatase (ALP), amylase (AMYL), pancreatic amylase (AMY-P), aspartate aminotransferase (AST), alanine aminotransferase (ALT), g-glutamyltransferase (GGT), lactate dehydrogenase (LDH), creatine kinase (CK), total bilirubin (BT), direct bilirubin (BD), phosphorus (P), calcium (Ca), magnesium (Mg), iron (Fe), sodium (Na), potassium (K), chloride (Cl), lipase, and hemolysis index (HI). The instrument was calibrated against appropriate proprietary reference standard materials and verified with the use of proprietary quality controls. Coagulation tests. All plasma aliquots were thawed at the same time. The routine laboratory tests were performed in duplicate immediately after thawing on the same instrument, ACL TOP (Instrumentation Laboratory, Milan, Italy), according to the manufacturer’s specifications and using HemosIL reagent system (proprietary reagents). The panel of tests included the following: aPTT, PT, fibrinogen (FIB) antithrombin (AT), protein C (PC), and protein S (PS). The instrument was calibrated against appropriate proprietary reference standard material and verified with the use of proprietary quality controls. A performance evaluation of the within-run precision of the automated coagulation analyzer ACL TOP in normal plasma showed coefficients of variation of 1.6% for aPTT.26
Statistical analysis The significance of the differences between samples was assessed using the paired Student t-test after verifying the normality by employing the D’Agostino-Pearson omnibus test. Because non-normal distributions were found for GLU, TG, CRP, urea, ALP, AMYL, AMY-P, AST, CK, BT, MG, NA, lipase, RBC, HGB, MCV, NEU, MPV, ESR, aPTT, PS, PC, and AT, results were assessed by the Wilcoxon rankedpairs test using licensed statistical software (GraphPad Prism version 5.01, La Jolla, CA, USA). Based on screening three different diagnostic blood specimen procedures in parallel, a p value < 0.017 was considered statistically significant, according to Bonferroni correction for
57
multiple comparisons. Mean % differences were determined according to the formula: mean % difference = [(mixing procedure x - mixing procedure y)/mixing procedure x] x 100. Finally, mean % differences from mixing procedures were compared with the current desirable quality specifications, derived from biological variation27,28,29 according to the formula: CVa < 0:5CVW jBAj < 0:25 (CVW 2 þ CVG 2 )0:5 where CVa is the percent analytical coefficient of variation, CVW is the within-subject biological variation, CVG is the between subject biological variation, and BA is the bias.
Results Significant statistical differences were found for: i) RBC and hematocrit when Procedure 1 was compared with Procedure 2; ii) ALT and ESR when Procedure 1 was compared with Procedure 3; iii) RBC, HCT and HI when Procedure 2 was compared with Procedure 3. Surprisingly, clinically significant differences were found only for Na when Procedure 1 was compared with Procedure 2, and Procedure 1 was compared with Procedure 3. The results of this investigation are shown in Table 1. No fibrin filaments or microclots were observed in samples from G1, G2, or G3.
Discussion Our results support the outcome from Parenmark and Landberg that the mixing of primary vacuum tubes is not mandatory.14 Parenmark and Landberg performed blood collection by venipuncture in vacuum tubes from 50 patients (10 or 20 patients in each group).14 To confirm their outcomes we decided to perform the blood collection from 100 volunteers for each group. When we compared the gold standard (Procedure 1) with no mixing (Procedure 3) for clinical chemistry results, only ALT showed a significant statistical difference ( p = 0.015). The gold standard mixing (Procedure 1) showed higher HI than the no mixing procedure (Procedure 3), although the HI that may generate a bias in results of ALT is higher than 700 according to claims of the company of the instrument(s) used in this study. Obviously an HI of 4.3 as in our study cannot be responsible for the statistically significant difference observed for ALT (Table 1). Moreover, no clinically significant difference was found when mean % differences from the gold standard (Procedure 1) versus no mixing (Procedure 3) were compared with the current desirable bias for quality specifications, derived from biological variation. As is widely known, a desirable bias is conventionally one of the three levels of quality specification, and it is the higher level of expectation in terms of quality, followed by optimal and minimal.27,28 The representation of results by mean % differences can appear obvious, since it is based on an algebraic calculation, but it may represent a useful tool in daily practice for quality managers of clinical laboratories.28–32 After applying this tool to the results of Parenmark and Landberg [14] (Table 2), we observed that our data (Table 1) are in partial accord with the above-mentioned study. The most important discrepancies were higher bias
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LIMA-OLIVEIRA ET AL.
Table 2. Mean % Difference Calculated from Parenmark and Landberg14 Mean % difference
Parameter Chemistry CRP AST LDH CK BT Fe Na K Hematology RBC HGB HCT MCV WBC PLT Coagulation PT APTT
Desirable bias (%)29
Instant mixing vs 5 min rest before mixing
Instant mixing vs no mix
21.8 5.4 4.3 11.5 11.4 8.8 0.3 1.8
0.0 2.4 2.8 0.7 - 0.7 - 0.8 1.4* 1.7
0.0 2.4 5.0* 0.7 - 0.1 0.8 1.4* 1.2
1.7 1.8 1.7 1.2 5.6 5.9
0.9 - 0.7 2.3** 0.11 2.3 4.3
0.9 0.0 2.3** 0.1 4.4 - 2.4
2.0 2.3
- 1.0 0.3
22.0* 0.7
*p < 0.05; **p value not communicated by authors.14 Bold mean % differences represent clinically significant variations, when compared with desirable bias.
than specified for PT, LDH, Na, and HCT when instant mixing was compared with 6 min rest in a supine position without mixing afterwards14 (the last procedure is equivalent to our Procedure 3). This is reasonable, because Parenmark and Landberg14 did not compare the no mixing procedure (equivalent to our Procedure 3) with a gold standard (e.g., procedure from the manufacturer’s datasheet, equivalent to our Procedure 1), but they used a horizontal mixing tray for 1 min (15 times inversion).14 As such, this procedure obviously increases the preanalytical variability and HI. It is also noteworthy that the horizontal mixing tray used by Parenmark and Landberg does not represent the manual mixing regularly used by phlebotomists in blood collection procedures. On the other hand, our results for hematology testing are in agreement with the previous experimental study from Lippi et al., where different mixing procedures (including no mixing) showed a percentage bias that did not exceed the quality specifications derived from biological variation.33 These authors compared the different mixing procedures with K2 EDTA primary tubes as follows: i) unmixed specimens versus inverting 6 times; and ii) unmixed specimens versus inverting 12 times. We are doubtful that different IVD manufacturers are providing equivalent products (as regards quality) to laboratory customers, since previous studies from our group have shown that vacuum tubes and dedicated syringes for blood collection from different producers should not be interchanged due to a lack of harmonization.34–38 Nevertheless, the present results (even sustained by the absence of micro-clots or fibrin filaments in all samples evaluated), suggest that the blood turbulence
generated by the standard vacuum pressure inside blood collection tubes39 is itself sufficient to grant a sufficient degree of solubilization, mixing, and stabilization of additives and blood during collection. Moreover, apparently incorrect vigorous mixing of the primary blood vacuum tubes does not appear to promote laboratory variability.40 In conclusion, our results show that failing to mix primary blood tubes after collection with a vacuum tube system does not introduce a bias in test results, or jeopardize the quality of testing. We conclude that mixing primary blood tubes is unnecessary. However, further studies should be planned to evaluate the impact of sample mixing/no-mixing when vacuum tubes are to be filled with blood is collected using open systems (e.g., syringe and needle).
Acknowledgments We are grateful to Monica Voi, Lara Salandini and Marina Saggiorno, medical technicians from the clinical chemistry section, hematology section, and coagulation section, respectively, for their skillful technical support. Special thanks to Ms. Sandra Meneghelli, quality manager.
Author Disclosure Statement No potential conflicts of interest relevant to this article were reported.
References 1. Geddes JT, Ahmed S, Pruetz BL, et al. SPIN: Development of sample-specific protein integrity numbers as an index of biospecimen quality. Biopreserv Biobank 2013; 11:25–32. 2. Kang HJ, Jeon SY, Park JS, et al. Identification of clinical biomarkers for pre-analytical quality control of blood samples. Biopreserv Biobank 2013;11:94–100. 3. Lippi G, Becan-McBride K, Behulova D, et al. Preanalytical quality improvement: In quality we trust. Clin Chem Lab Med 2013;51:229–241. 4. Simundic AM, Lippi G. Preanalytical phase—A continuous challenge for laboratory professionals. Biochem Med 2012; 22:145–149. 5. Simundic AM, Bilic-Zulle L, Nikolac N, et al. The quality of the extra-analytical phase of laboratory practice in some developing European countries and Mexico. A multicentric study. Clin Chem Lab Med 2011;49:215–228. 6. Lippi G, Chance JJ, Church S, et al. Preanalytical quality improvement: From dream to reality. Clin Chem Lab Med 2011;49:1113–1126. 7. Lippi G, Simundic A-M. Total quality in laboratory diagnostics. It’s time to think outside the box. Biochem Med 2010;20:5–8. 8. Lippi G, Salvagno GL, Montagnana M, et al. Quality standards for sample collection in coagulation testing. Semin Thromb Hemost 2012;38:565–575. 9. Lima-Oliveira G, Guidi GC, Salvagno GL, et al. Is phlebotomy part of the dark side in the clinical laboratory struggle for quality? Lab Med 2012;43:17–21. 10. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Impact of the phlebotomy training based on CLSI/NCCLS H03-A6. Procedures for the collection of diagnostic blood specimens by venipuncture. Biochem Med 2012;22:342–351. 11. Nikolac N, Supak-Smolcic V, Simundic AM, et al. Croatian Society of Medical Biochemistry and Laboratory Medicine:
MIXING OF VACUUM TUBES
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National recommendations for venous blood sampling. Biochem Med 2013;23:242–254. Clinical Laboratory Standards Institute. Procedures for the Handling and Processing of Blood Specimens for Common Laboratory Tests. CLSI H18-A4 document. 4th ed. Wayne, PA: Clinical Laboratory Standards Institute; 2010. Clinical Laboratory Standards Institute. Collection, Transport, and Processing of Blood Specimens for Testing Plasma-Based Coagulation Assays and Molecular Hemostasis Assays. CLSI H21-A5 document. 5th ed. Wayne, PA: Clinical Laboratory Standards Institute; 2008. Parenmark A, Landberg E. To mix or not to mix venous blood samples collected in vacuum tubes? Clin Chem Lab Med 2011;49:2061–2063. International Organization for Standardization. Medical Laboratories—Particular Requirements for Quality and Competence ISO Document 15189. 2nd ed. Geneva, Switzerland: International Organization for Standardization; 2007. CLSI. Procedures for the collection of diagnostic blood specimens by venipuncture. NCCLS H3-A6. 6 ed2007. Guder WG, Narayanan S, Wisser H, et al. Diagnostic Samples: From the patient to the Laboratory: The Impact of Preanalytical Variables on the Quality of Laboratory Results. 4 ed: Wiley-Blackwell; 2009. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Transillumination: A new tool to eliminate the impact of venous stasis during the procedure for the collection of diagnostic blood specimens for routine haematological testing. Int J Lab Hematol 2011;33:457–462. Lima-Oliveira G, Salvagno GL, Lippi G, et al. Elimination of the venous stasis error for routine coagulation testing by transillumination. Clin Chim Acta 2011;412:1482–1484. Lima-Oliveira G, Lippi G, Salvagno GL, et al. New ways to deal with known preanalytical issues: Use of transilluminator instead of tourniquet for easing vein access and eliminating stasis on clinical biochemistry. Biochem Med 2011;21:152–159. CLSI. Procedures for the Handling and Processing of Blood Specimens for Common Laboratory Tests. NCCLS H18-A4. 4 ed, 2010. CLSI. Collection, Transport, and Processing of Blood Specimens for Testing Plasma-Based Coagulation Assays and Molecular Hemostasis Assays. NCCLS H21-A5. 5 ed2008. Terumo. Venosafe Plastic Tubes: Coagulation—VF054SBCS07. Terumo Europe N.V.; 2012. Available from: http://www.terumo-europe.com/laboratory/products/vf_054 sbcs07.php. Accessed 2 April 2013. Terumo. Venosafe Plastic Tubes: Plasma Gel—VF054SAHL. Terumo Europe N.V.; 2013. Available from: http://www.terumo-europe.com/laboratory/products/vf_054 sahl.php. Accessed 2 April 2013. Terumo. Venosafe Plastic Tubes: Hematology—VF053SDK. Terumo Europe N.V.; 2012. Available from: http://www.terumo-europe.com/laboratory/products/vf_053 sdk.php. Accessed 2 April 2013. Milos M, Herak D, Kuric L, et al. Evaluation and performance characteristics of the coagulation system: ACL TOP analyzer—HemosIL reagents. Int J Lab Hematol 2009;31: 26–35.
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27. Hyltoft Petersen P, Fraser CG. Strategies to set global analytical quality specifications in laboratory medicine: 10 years on from Stockholm consensus conference. Accred Qual Assur 2010;15:323–330. 28. Ricos C, Alvarez V, Cava F, et al. Current databases on biological variation: pros, cons and progress. Scand J Clin Lab Invest 1999;59:491–500. 29. Westgard J. Biological Variation Database Specifications. 2010 [updated january 2012. Available from: http://www .westgard.com/biodatabase1.htm. Accessed 2 April 2013. 30. Ricos C, Cava F, Garcia-Lario JV, et al. The reference change value: A proposal to interpret laboratory reports in serial testing based on biological variation. Scand J Clin Lab Invest 2004;64:175–184. 31. Cembrowski GS, Tran DV, Higgins TN. The use of serial patient blood gas, electrolyte and glucose results to derive biologic variation: a new tool to assess the acceptability of intensive care unit testing. Clin Chem Lab Med 2010;48: 1447–1454. 32. Plebani M, Lippi G. Biological variation and reference change values: An essential piece of the puzzle of laboratory testing. Clin Chem Lab Med 2012;50:189–190. 33. Lippi G, Salvagno GL, Montagnana M, et al. Evaluation of different mixing procedures for K2 EDTA primary samples on hematological testing. Lab Med 2007;38:723–725. 34. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Different manufacturers of syringes: A new source of variability in blood gas, acid-base balance and related laboratory test? Clin Biochem 2012;45:683–687. 35. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Preanalytical management: Serum vacuum tubes validation for routine clinical chemistry. Biochem Med 2012;22:180–186. 36. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Sodium citrate vacuum tubes validation: Preventing preanalytical variability in routine coagulation testing. Blood Coagul Fibrinolysis 2013;24:252–255. 37. Lima-Oliveira G, Lippi G, Salvagno GL, et al. K3EDTA vacuum tubes validation for routine hematological testing. ISRN Hematology 2012;2012:875357. 38. Lima-Oliveira G, Salvagno GL, Lippi G, et al. Quality management of preanalytical phase: Impact of lithium heparin vacuum tubes changes on clinical chemistry. Accred Qual Assur 2013;18:429–434. 39. International Organization for Standardization. Single-use containers for venous blood specimen collection. ISO 6710 document. Geneva, Switzerland: International Organization for Standardization; 1995. 40. Lima-Oliveira G, Lippi G, Salvagno GL, et al. Effects of vigorous mixing of blood vacuum tubes on laboratory test results. Clin Biochem 2013;46:250–254.
Address correspondence to: Gabriel Lima-Oliveira, MsC. Av. Pref. Lothario Meissner 632 Jardim Botanico Curitiba Parana 80210-170 Brazil E-mail: [email protected]
Processing of Diagnostic Blood Specimens: Is It Really Necessary to Mix Primary Blood Tubes after Collection with Evacuated Tube System? Gabriel Lima-Oliveira,1,2 Giuseppe Lippi,3 Gian Luca Salvagno,1 Giorgio Brocco,1 Stefania Gaino,1 Francesco Dima,1 Waldemar Volaski,2 Fabiane Gomes Rego,2 Geraldo Picheth,2 and Gian Cesare Guidi1,2
Background: The preanalytical phase is considered the most vulnerable phase in biopreservation, biobanking, and laboratory diagnostics. Accurate mixing after blood collection is claimed to be important and recommended by the manufacturers. Objective: To evaluate whether it is really necessary to mix the primary blood tubes immediately after blood collection by means of evacuated tube systems. Material and Methods: Blood from 300 outpatients was equally and randomly divided into three groups: G1, sodium citrate vacuum tubes; G2, lithium heparin vacuum tubes; and G3, K2EDTA vacuum tubes. All vacuum tubes were processed using three different procedures. Procedure 1: Gold Standard (P1): All specimens mixed gently and carefully by inverting five times as recommended; Procedure 2: Rest time (P2): All specimens remained 5 min in the upright position, followed by gentle careful mixing by inverting five times; Procedure 3: No mix (P3): All specimens were left in upright position without mixing afterwards. The influence of the primary mixing tube procedure was evaluated for clinical chemistry, hematology, and coagulation parameters by paired t-test. The bias from the mixing procedure was also compared with quality specifications derived from biological variation. Results: Significant differences ( p < 0.017) were found for: i) red blood cell count and hematocrit when P1 was compared with P2; ii) alanine aminotransferase and erythrocyte sedimentation rate when P1 was compared with P3; iii) red blood cell count, hematocrit, and hemolysis index when P2 was compared with P3. Surprisingly, clinically significant differences were found only for sodium when P1 was compared with P2, and P1 was compared with P3. No fibrin filaments or microclots were observed in any samples. Conclusion: Primary blood tubes mixing after collection with evacuated tube system appears to be unnecessary.
key factors in maintaining quality of biospecimens.2 Presently, quality managers focus their attention on the preanalytical phase.3–11 Nevertheless, the manually intensive activities of the preanalytical phase are still considered the most vulnerable steps in biopreservation, biobanking, and laboratory diagnostics. In this field of activity, accurate mixing of tubes with anticoagulant or clot activator additives after blood collection is considered essential and is widely recommended by manufacturers’ datasheets.12,13 Parenmark and Landberg14 recently released a convincing paradigm about the mixing procedure for diagnostic blood specimens. They showed that: i) mixing blood samples immediately after collection may be not mandatory for all types of tubes; and ii) instant mixing may produce spurious
Introduction
B
iomarkers are measurable characteristics represented by proteins, enzymes, ions, and/or macromolecules and cells, which can be used to screen, diagnose, and monitor diseases. It is well known that the process of finding and validating suitable biomarkers is important for clinical research and disease management. Researchers, physicians, and laboratory managers looking for biomarkers may save a large amount of time and money when samples from biobanks are obtained through use of a quality management system.1 Indeed, sample collection and processing are essential steps for biopreservation and biobanking, so that the control of standardized preanalytical conditions are 1
Laboratory of Clinical Biochemistry, Department of Life and Reproduction Sciences, University of Verona, Verona, Italy. Post-Graduate Program of Pharmaceutical Sciences, Department of Medical Pathology Federal University of Parana, Curitiba, Parana, Brazil. 3 Pathology and Laboratory Medicine, Clinical Chemistry and Hematology Laboratory, Academic Hospital of Parma, Parma, Italy. 2
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LIMA-OLIVEIRA ET AL.
hemolysis and thereby introduce a bias for those parameters that are most susceptible to erythrocyte injury.14 The authors also concluded that their study was limited, and results could only be considered as preliminary findings. The main drawback in the outcome of this study was that mixing was performed by putting the tube on a horizontal mixing tray for 1 min (inverting 15 times). This procedure is however different from the manual mixing procedure that is conventionally used by phlebotomists. On this basis alone, the conclusion of increased hemolysis due to immediate mixing could be misleading and cannot be generalized; iii) immediate mixing resulted in a higher plasma hemolysis index in citrate tubes than in heparin tubes, a rather odd finding. Despite this, there was no significant difference between immediate mixing versus 5 min rest before mixing for prothrombin time (PT) and activated partial thromboplastin time (aPTT) results (nor any significant difference in potassium [K] and lactate dehydrogenase [LDH] in heparin tubes). Therefore, this single, though relevant, study provides scant support for an adverse impact of immediate mixing. Relevant to this point, the ISO 15189 standard states that necessary improvements and potential sources of nonconformities, either technical or concerning the quality management system, shall be identified and all laboratory processes shall be validated.15 The aim of the present study was to evaluate whether it is really necessary to mix the primary blood tubes immediately after blood collection using evacuated tube systems.
Europe, Leuven, Belgium) directly into three vacuum tubes, as follows: G1) three 3.6 mL vacuum tubes containing 0.4 mL of buffered sodium citrate (9NC) 0.109 mol/L: 3.2 W/V% (Terumo Europe, Leuven, Belgium); G2) three 3.5 mL vacuum tubes with 52.5 USP U of lithium heparin and gel separator (Terumo Europe); and G3) three 3.0 mL vacuum tubes containing 5.9 mg K2EDTA (Terumo Europe). To eliminate any potential interference due to either the contact phase or the tissue factor, approximately 2 mL of blood was preliminarily collected in a discard tube without additive (Vacuette, Greiner Bio-One GmbH, Kremsmu¨nster, Austria). Blood collection was standardized, including the use of needles and vacuum tubes of the same lot.
Materials and Methods
Procedure 3—No mixing. All diagnostic blood specimens were left in the upright position (i.e., tube stopper at the top), without mixing afterwards. Centrifugation. All samples were centrifuged immediately within 10 min after blood collection as follows: i) sodium citrate vacuum tubes at 1500 g for 15 min at room temperature; and ii) lithium heparin vacuum tubes at 1200 g for 10 min at room temperature. All plasma citrate samples were separated, and because shortening of aPTT in thawed samples may also be caused by residual platelet debris in the thawed sample, a re-centrifugation was done and the sample was then aliquoted. This is the usual practice, to ensure platelet-free citrate samples for freezing. Diagnostic blood specimens storage. All plasma samples, both from citrate and lithium heparin-vacuum tubes were stored in aliquots and kept frozen at - 70C until measurement (22 days after the first blood collection). Samples did not show any signs of hemolysis by visual inspection. No specimen was discarded due to unsatisfactory attempts, difficult venous access, missing veins, manifest hemolysis, or lipemia.
Study design Three hundred adult outpatients of both sexes were invited by the Laboratory of Clinical Biochemistry, (132 males, 168 females, mean age – SD 41 – 3), Department of Life and Reproduction Sciences, University of Verona, Italy, to participate in the study, by allowing three extra tubes to be drawn (*12 mL of additional blood over the normal amount collected by the phlebotomist for routine analysis). These volunteers were equally and randomly divided into three groups (Parenmark protocol)14 as follows: i) G1 sodium citrate vacuum tubes; ii) G2 lithium heparin vacuum tubes; and iii) G3 K2EDTA vacuum tubes. Each volunteer agreed through informed consent to be enrolled in this study, which was also carried out in agreement with the Declaration of Helsinki and under the terms of all relevant local regulations.
Collection of diagnostic blood specimens The collection of blood specimens was performed from 8:00 to 9:00 AM, over a period of 3 weeks by a single expert phlebotomist, according to the recommendations of the Clinical Laboratory Standard Institute (CLSI).16 All volunteers, after 12-hours fasting, were maintained in a sitting position for 15 min prior to phlebotomy in order to eliminate possible interference of blood distribution due to posture.17 After this time frame, a radial vein was located on the right forearm by a subcutaneous tissue transilluminator device (Venosco´pio IV plus, Duan do Brasil, Sao Paulo, Brazil) without tourniquet, to prevent interference from venous stasis,18–20 and approximately 12 mL of blood was collected by venipuncture with a 20-gauge straight needle (Terumo
Processing of diagnostic blood specimens Primary blood tube mixing. All vacuum tubes from G1, G2, and G3 were processed through three different procedures. Procedure 1—Gold Standard.21,22 All diagnostic blood spec-
imens were mixed gently and carefully by inverting five times, —immediately after blood collection—as recommended by the manufacturer.23–25 Procedure 2—Rest time. All diagnostic blood specimens after collection had a 5 min rest in the upright position (i.e., tube stopper at the top), followed by gentle and careful mixing by inverting five times .
Laboratory testing All laboratory tests were performed in duplicate to minimize the analytical coefficient of variation (CVa), so the CVa from analyses in duplicate is smaller than the CVa from internal quality control shown in Table 1. Assessment of hematological parameters. All samples were processed for routine hematological testing immediately after collection ( < 15 min) on the same Sysmex XE2100D, Automated Hematology Analyzer (Sysmex Corporation, Kobe, Japan). The parameters tested included
55
GLU* COL** HDL** TG* TP** ALB** CRP* UREA* CRE** UA** ALP* AMYL* AMY-P* AST* ALT** GGT** LDH** CK* BT* BD** P** Ca** Mg* Fe** Na* K** Cl** Lipase* HI**
Tests
(mmol/L) (mmol/L) (mmol/L) (mmol/L) (g/L) (g/L) (mg/L) (mmol/L) (mmol/L) (mmol/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mkat/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mmol/L) (mkat/L) NA
(Units)
2.2 4.0 5.2 10.7 1.2 1.3 21.8 5.5 3.8 4.9 6.4 7.4 8.0 5.4 12.0 10.8 4.3 11.5 11.4 14.2 3.2 0.8 1.8 8.8 0.3 1.8 0.5 10.1 NA
1.5 1.8 4.3 2.0 1.2 1.2 2.2 2.6 2.5 1.0 2.7 0.8 2.2 1.2 1.3 1.6 1.5 3.3 2.2 1.8 3.0 0.7 1.2 2.6 1.0 1.5 1.0 2.8 NA
6.10 [5.33 – 8.71] 4.64 – 0.97 1.38 – 0.30 1.20 [0.85 – 1.60] 73.2 – 5.87 41.5 – 3.37 2.20 [1.40 – 3.70] 6.82 [5.96 – 8.32] 83.1 – 17.7 0.34 – 0.08 1.10 [0.90 – 1.34] 1.02 [0.80 – 1.29] 0.37 [0.34 – 0.48] 0.31 [0.27 – 0.46] 0.31 – 0.09 0.85 – 0.18 5.66 – 0.26 1.41 [1.12 – 1.97] 9.40 [7.70 – 14.7] 3.93 – 2.05 0.98 – 0.18 2.36 – 0.08 0.78 [0.76 – 0.84] 79.6 – 25.9 141 [139 – 142] 4.40 – 0.52 101 – 2.66 0.54 [0.44 – 0.83] 4.3 – 1.8
Procedure 1 Gold Standard
Procedure 3 No mix
6.16 [5.33 – 8.55] 4.64 – 0.95 1.38 – 0.30 1.22 [0.85 – 1.57] 73.2 – 5.72 41.5 – 3.24 2.20 [1.30 – 3.70] 6.68 [6.07 – 8.42] 81.3 – 17.7 0.34 – 0.08 1.10 [0.90 – 1.34] 1.00 [0.78 – 1.29] 0.36 [0.34 – 0.46] 0.31 [0.29 – 0.44] 0.30 – 0.10 0.85 – 0.18 5.61 – 0.26 1.39 [1.12 – 1.97] 9.75 [7.70 – 14.0] 4.10 – 2.05 0.98 – 0.17 2.36 – 0.09 0.78 [0.75 – 0.85] 79.0 – 25.8 141 [139 – 142] 4.39 – 0.53 101 – 2.63 0.54 [0.44 – 0.83] 4.0 – 2.8
Clinical chemistry 6.10 [5.22 – 8.49] 4.64 – 0.94 1.38 – 0.30 1.18 [0.85 – 1.57] 72.8 – 6.13 41.2 – 3.36 2.20 [1.40 – 3.70] 6.78 [6.10 – 8.71] 82.2 – 16.8 0.34 – 0.08 1.10 [0.88 – 1.36] 1.00 [0.78 – 1.28] 0.36 [0.34 – 0.46] 0.31 [0.29 – 0.42] 0.30 – 0.09 0.86 – 0.18 5.64 – 0.26 1.41 [1.16 – 1.96] 9.58 [7.70 – 13.7] 3.93 – 2.05 0.97 – 0.16 2.36 – 0.10 0.77 [0.74 – 0.85] 79.0 – 25.1 140 [139 – 141] 4.39 – 0.52 101 – 2.75 0.54 [0.44 – 0.85] 4.2 – 2.0
Procedure 2 Rest time
vs
vs
vs
Procedure 2 Rest time
0.0 0.0 0.0 1.7 0.6 0.7 0.0 0.6 1.1 0.0 0.0 2.0 2.7 0.0 3.2 - 1.2 0.4 0.0 - 1.9 0.0 1.0 0.0 1.3 0.8 0.7 0.2 0.0 0.0 2.3
mean % difference
0.1255 0.3425 1.000 0.0294 0.0905 0.0863 0.0975 0.2377 0.2189 0.8534 0.2153 0.2759 0.1096 0.3026 0.1036 0.4818 0.8063 0.4246 0.0985 0.3855 0.2001 0.5161 0.9045 0.2048 0.1121 0.6514 0.1429 0.8211 0.6669
P value
1.0 0.0 0.0 1.6 0.0 0.0 0.0 - 2.1 - 2.2 0.0 0.0 - 2.0 - 2.8 0.0 - 3.3 0.0 - 0.9 - 1.4 3.6 4.2 0.0 0.0 0.0 - 0.8 0.0 - 0.2 0.0 0.0 - 7.5
mean % difference
0.7747 0.4731 0.1716 0.4032 0.7746 0.6001 0.9052 0.6468 0.1953 0.4818 0.6675 1.000 0.0708 0.6444 0.0150 0.7899 0.1851 0.7432 0.5987 0.7551 0.5463 0.4909 0.9430 0.0465 0.3363 0.3728 0.1628 0.5877 0.6012
P value
- 1.0 0.0 0.0 - 3.4 - 0.6 - 0.7 0.0 1.5 1.1 0.0 0.0 0.0 0.0 0.0 0.0 1.2 0.5 1.4 - 1.8 - 4.3 - 1.0 0.0 - 1.3 0.0 20.7 0.0 0.0 0.0 4.8
mean % difference
(continued)
0.0498 0.8233 0.1716 0.2350 0.1115 0.1863 0.1589 0.6871 0.5725 0.6669 0.3778 0.2555 0.7768 0.4070 0.1868 0.1036 0.1757 0.9638 0.2005 0.3306 0.0841 0.8097 0.8864 1.0000 0.1161 0.9534 0.6720 0.5297 0.8072
P value
Procedure 2—Rest time Procedure 3—No mix Procedure 3—No mix
Procedure 1 Gold Standard
Procedure 1 Gold Standard
Effect of Three Different Procedures of Primary Blood Vacuum Tubes Mixing in Laboratory Testing
Desirable CVa Bias (%)29 (%)
Table 1.
56
1.3 1.2 3.6 7.0 2.0 3.0 NA
4.64 [4.34 – 4.74] 137 [129 – 148] 43.5 – 4.50 96.1 [93.5 – 98.1] 13.1 – 0.63 57.6 – 3.70 6.14 – 1.36 2.96 [2.50 – 3.92] 1.99 – 0.47 0.41 – 0.14 0.20 – 0.03 0.03 – 0.01 0.13 – 0.01 233 – 13.3 9.05 [8.70 – 9.70] 24.0 [17.2 – 34.8]
31.4 – 10.7 38.9 [35.7 – 44.2] 3.08 – 0.15 34.0 [29.5 – 45.0] 46.0 [39.0 – 52.0] 89.0 [82.0 – 102.0] 2.0 [2.0 – 3.0]
31.1 – 10.5 38.6 [36.0 – 45.4] 3.13 – 0.16 34.0 [29.0 – 46.0] 46.0 [38.0 – 55.0] 89.0 [83.0 – 102.0] 3.0 [2.0 – 4.0]
Procedure 2 Rest time
4.67 [4.36 – 4.87] 138 [129 – 150] 43.9 – 1.02 96.2 [93.2 – 98.1] 13.1 – 0.63 58.9 – 3.88 5.97 – 1.36 3.01 [2.50 – 3.87] 1.95 – 0.47 0.41 – 0.15 0.20 – 0.03 0.03 – 0.01 0.13 – 0.01 232 – 13.5 9.10 [8.62 – 9.48] 24.5 [17.2 – 38.0]
Procedure 1 Gold Standard
31.6 – 11.0 38.5 [36.6 – 45.6] 3.10 – 0.16 34.0 [29.0 – 44.5] 46.0 [35.0 – 52.0] 87.0 [81.0 – 102.0] 4.0 [2.0 – 4.0]
Coagulation
4.68 [4.35 – 4.80] 138 [130 – 149] 43.8 – 1.01 96.2 [93.4 – 98.1] 13.1 – 0.63 58.6 – 3.66 6.10 – 1.35 2.99 [2.42 – 3.87] 1.96 – 0.52 0.41 – 0.12 0.21 – 0.03 0.03 – 0.01 0.13 – 0.01 231 – 12.6 9.05 [8.72 – 9.48] 24.0 [18.0 – 35.0]
Hematology
Procedure 3 No mix
vs
vs
vs
Procedure 2 Rest time
- 1.0 - 0.8 1.6 0.0 0.0 0.0 33.3
0.6 0.7 0.9 0.1 0.0 2.2 - 2.8 1.7 - 2.0 0.0 0.0 0.0 0.0 - 0.4 0.6 2.0
mean % difference
0.0790 0.6493 0.2931 0.1195 0.0724 0.7315 0.3360
0.0037 0.2115 0.0012 0.5648 0.8037 0.2835 0.1442 0.9358 0.0545 0.4540 0.7894 0.2674 0.2538 0.6843 0.2756 0.0676
P value
1.6 - 0.3 - 1.0 0.0 0.0 - 2.3 25
0.2 0.0 - 0.2 0.0 0.0 - 0.5 2.1 - 0.7 0.5 0.0 4.8 0.0 0.0 - 0.4 - 0.6 - 2.1
mean % difference
0.0619 0.9224 0.4396 0.0315 0.1006 0.0804 0.1839
0.5374 0.0664 0.4540 0.3194 0.2044 0.8755 0.2107 0.7627 0.6736 0.7524 0.2407 0.0375 0.8098 0.6653 0.1311 0.0033
P value
- 0.6 1.0 - 0.6 0.0 0.0 2.2 - 100
- 0.9 - 0.7 - 0.7 - 0.1 0.0 - 1.7 0.6 - 1.0 1.5 0.0 - 5.0 0.0 0.0 0.9 0.0 0.0
mean % difference
0.4059 0.2872 0.7732 0.6444 0.2548 0.0697 0.0106
0.0068 0.3210 0.0051 0.1232 0.3590 0.3143 0.3869 0.6011 0.2055 0.7250 0.2813 0.3590 0.3665 0.2877 0.5752 0.7272
P value
Procedure 2—Rest time Procedure 3—No mix Procedure 3—No mix
Procedure 1 Gold Standard
Procedure 1 Gold Standard
*Non-normal distribution; the values are presented as median [interquartile range]; P value represents the significance by Wilcoxon ranked-pairs test. **Normal distribution; the values are presented as mean – standard deviation; P value represents the significance by paired Student’s t-test. The bold p values are statistically significant ( p < 0.017),29 according to Bonferroni correction for multiple comparisons. Mean % difference in bold was higher than the desirable bias. CVa (%): Analytical coefficient (within-run precision), by internal quality control on the respective instrument.
2.0 2.3 4.8 15.9 13.9 4.0 NA
(s) (s) (g/L) (%) (%) (%) NA
PT** aPTT* FIB** PS* PC* AT* HI*
1.5 1.0 1.5 1.0 2.0 6.5 3.0 8.0 8.0 9.8 15.5 9.7 7.8 4.0 1.2 8.9
Desirable CVa Bias (%)29 (%)
(10 / L) 1.7 (g/L) 1.8 (%) 1.7 (fL) 1.2 (%) 1.7 (109/ L) 7.8 (109/ L) 5.6 9.1 (106/ L) (106/ L) 7.4 (106/ L) 13.2 (106/ L) 19.8 15.4 (106/ L) (106/ L) NA 5.9 (109/ L) (fL) 2.3 mm/h NA
12
(Units)
RBC* HGB* HCT** MCV* RDW** Retic** WBC** NEU* LYMP** MONO** EOS** BASO** LUC** PLT** MPV* ESR*
Tests
Table 1. (Continued)
MIXING OF VACUUM TUBES
red blood cell count (RBC), hemoglobin (HGB), hematocrit (HCT), mean corpuscular volume (MCV), RBC distribution width (RDW), reticulocytes (Retic), white blood cells (WBC) count and differential, including neutrophils (NEU), lymphocytes (LYMP), monocytes (MONO), eosinophils (EOS), and basophils (BASO), platelet count (PLT), and mean platelet volume (MPV). The instrument had been previously calibrated against appropriate proprietary reference standard material and verified with the use of proprietary controls. The erythrocyte sedimentation rate (ESR) was measured in duplicate in the same TEST 1 YDL (Alifax, Padova, Italy) immediately after routine hematological testing on Sysmex XE-2100D. Clinical chemistry tests. All frozen lithium heparin aliquots were thawed at the same time. The routine clinical chemistry tests were performed on the same instrument Cobas 6000 < c501 > module (Roche Diagnostics GmbH, Penzberg, Germany), according to the manufacturer’s specifications and using proprietary reagents. The panel of tests included the following: glucose (GLU), total cholesterol (COL), high-density lipoprotein cholesterol (HDL), triglycerides (TG), total protein (TP), albumin (ALB), Creactive protein (CRP), urea, creatinine (CRE), uric acid (UA), alkaline phosphatase (ALP), amylase (AMYL), pancreatic amylase (AMY-P), aspartate aminotransferase (AST), alanine aminotransferase (ALT), g-glutamyltransferase (GGT), lactate dehydrogenase (LDH), creatine kinase (CK), total bilirubin (BT), direct bilirubin (BD), phosphorus (P), calcium (Ca), magnesium (Mg), iron (Fe), sodium (Na), potassium (K), chloride (Cl), lipase, and hemolysis index (HI). The instrument was calibrated against appropriate proprietary reference standard materials and verified with the use of proprietary quality controls. Coagulation tests. All plasma aliquots were thawed at the same time. The routine laboratory tests were performed in duplicate immediately after thawing on the same instrument, ACL TOP (Instrumentation Laboratory, Milan, Italy), according to the manufacturer’s specifications and using HemosIL reagent system (proprietary reagents). The panel of tests included the following: aPTT, PT, fibrinogen (FIB) antithrombin (AT), protein C (PC), and protein S (PS). The instrument was calibrated against appropriate proprietary reference standard material and verified with the use of proprietary quality controls. A performance evaluation of the within-run precision of the automated coagulation analyzer ACL TOP in normal plasma showed coefficients of variation of 1.6% for aPTT.26
Statistical analysis The significance of the differences between samples was assessed using the paired Student t-test after verifying the normality by employing the D’Agostino-Pearson omnibus test. Because non-normal distributions were found for GLU, TG, CRP, urea, ALP, AMYL, AMY-P, AST, CK, BT, MG, NA, lipase, RBC, HGB, MCV, NEU, MPV, ESR, aPTT, PS, PC, and AT, results were assessed by the Wilcoxon rankedpairs test using licensed statistical software (GraphPad Prism version 5.01, La Jolla, CA, USA). Based on screening three different diagnostic blood specimen procedures in parallel, a p value < 0.017 was considered statistically significant, according to Bonferroni correction for
57
multiple comparisons. Mean % differences were determined according to the formula: mean % difference = [(mixing procedure x - mixing procedure y)/mixing procedure x] x 100. Finally, mean % differences from mixing procedures were compared with the current desirable quality specifications, derived from biological variation27,28,29 according to the formula: CVa < 0:5CVW jBAj < 0:25 (CVW 2 þ CVG 2 )0:5 where CVa is the percent analytical coefficient of variation, CVW is the within-subject biological variation, CVG is the between subject biological variation, and BA is the bias.
Results Significant statistical differences were found for: i) RBC and hematocrit when Procedure 1 was compared with Procedure 2; ii) ALT and ESR when Procedure 1 was compared with Procedure 3; iii) RBC, HCT and HI when Procedure 2 was compared with Procedure 3. Surprisingly, clinically significant differences were found only for Na when Procedure 1 was compared with Procedure 2, and Procedure 1 was compared with Procedure 3. The results of this investigation are shown in Table 1. No fibrin filaments or microclots were observed in samples from G1, G2, or G3.
Discussion Our results support the outcome from Parenmark and Landberg that the mixing of primary vacuum tubes is not mandatory.14 Parenmark and Landberg performed blood collection by venipuncture in vacuum tubes from 50 patients (10 or 20 patients in each group).14 To confirm their outcomes we decided to perform the blood collection from 100 volunteers for each group. When we compared the gold standard (Procedure 1) with no mixing (Procedure 3) for clinical chemistry results, only ALT showed a significant statistical difference ( p = 0.015). The gold standard mixing (Procedure 1) showed higher HI than the no mixing procedure (Procedure 3), although the HI that may generate a bias in results of ALT is higher than 700 according to claims of the company of the instrument(s) used in this study. Obviously an HI of 4.3 as in our study cannot be responsible for the statistically significant difference observed for ALT (Table 1). Moreover, no clinically significant difference was found when mean % differences from the gold standard (Procedure 1) versus no mixing (Procedure 3) were compared with the current desirable bias for quality specifications, derived from biological variation. As is widely known, a desirable bias is conventionally one of the three levels of quality specification, and it is the higher level of expectation in terms of quality, followed by optimal and minimal.27,28 The representation of results by mean % differences can appear obvious, since it is based on an algebraic calculation, but it may represent a useful tool in daily practice for quality managers of clinical laboratories.28–32 After applying this tool to the results of Parenmark and Landberg [14] (Table 2), we observed that our data (Table 1) are in partial accord with the above-mentioned study. The most important discrepancies were higher bias
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LIMA-OLIVEIRA ET AL.
Table 2. Mean % Difference Calculated from Parenmark and Landberg14 Mean % difference
Parameter Chemistry CRP AST LDH CK BT Fe Na K Hematology RBC HGB HCT MCV WBC PLT Coagulation PT APTT
Desirable bias (%)29
Instant mixing vs 5 min rest before mixing
Instant mixing vs no mix
21.8 5.4 4.3 11.5 11.4 8.8 0.3 1.8
0.0 2.4 2.8 0.7 - 0.7 - 0.8 1.4* 1.7
0.0 2.4 5.0* 0.7 - 0.1 0.8 1.4* 1.2
1.7 1.8 1.7 1.2 5.6 5.9
0.9 - 0.7 2.3** 0.11 2.3 4.3
0.9 0.0 2.3** 0.1 4.4 - 2.4
2.0 2.3
- 1.0 0.3
22.0* 0.7
*p < 0.05; **p value not communicated by authors.14 Bold mean % differences represent clinically significant variations, when compared with desirable bias.
than specified for PT, LDH, Na, and HCT when instant mixing was compared with 6 min rest in a supine position without mixing afterwards14 (the last procedure is equivalent to our Procedure 3). This is reasonable, because Parenmark and Landberg14 did not compare the no mixing procedure (equivalent to our Procedure 3) with a gold standard (e.g., procedure from the manufacturer’s datasheet, equivalent to our Procedure 1), but they used a horizontal mixing tray for 1 min (15 times inversion).14 As such, this procedure obviously increases the preanalytical variability and HI. It is also noteworthy that the horizontal mixing tray used by Parenmark and Landberg does not represent the manual mixing regularly used by phlebotomists in blood collection procedures. On the other hand, our results for hematology testing are in agreement with the previous experimental study from Lippi et al., where different mixing procedures (including no mixing) showed a percentage bias that did not exceed the quality specifications derived from biological variation.33 These authors compared the different mixing procedures with K2 EDTA primary tubes as follows: i) unmixed specimens versus inverting 6 times; and ii) unmixed specimens versus inverting 12 times. We are doubtful that different IVD manufacturers are providing equivalent products (as regards quality) to laboratory customers, since previous studies from our group have shown that vacuum tubes and dedicated syringes for blood collection from different producers should not be interchanged due to a lack of harmonization.34–38 Nevertheless, the present results (even sustained by the absence of micro-clots or fibrin filaments in all samples evaluated), suggest that the blood turbulence
generated by the standard vacuum pressure inside blood collection tubes39 is itself sufficient to grant a sufficient degree of solubilization, mixing, and stabilization of additives and blood during collection. Moreover, apparently incorrect vigorous mixing of the primary blood vacuum tubes does not appear to promote laboratory variability.40 In conclusion, our results show that failing to mix primary blood tubes after collection with a vacuum tube system does not introduce a bias in test results, or jeopardize the quality of testing. We conclude that mixing primary blood tubes is unnecessary. However, further studies should be planned to evaluate the impact of sample mixing/no-mixing when vacuum tubes are to be filled with blood is collected using open systems (e.g., syringe and needle).
Acknowledgments We are grateful to Monica Voi, Lara Salandini and Marina Saggiorno, medical technicians from the clinical chemistry section, hematology section, and coagulation section, respectively, for their skillful technical support. Special thanks to Ms. Sandra Meneghelli, quality manager.
Author Disclosure Statement No potential conflicts of interest relevant to this article were reported.
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Address correspondence to: Gabriel Lima-Oliveira, MsC. Av. Pref. Lothario Meissner 632 Jardim Botanico Curitiba Parana 80210-170 Brazil E-mail: [email protected]
