with 50 g/L OLZ added. As can be seen in Fig

Clinical Chemistry 49, No. 12, 2003
with 50 ␮g/L OLZ added. As can be seen in Fig. 1D,
extracts from protein precipitation and solid-phase methods showed a substantial reduction in instrument response compared with mobile phase. In contrast, the ethyl
acetate liquid–liquid preparation used in our assay suppressed the signal of the analytes by ⬍3%.
This assay, because of the high simplicity of the extraction procedure, the short duration of the analysis, the
small amount of starting sample needed, and its high
selectivity, is being used routinely by our research group
to monitor plasma OLZ concentrations in schizophrenic
patients. In fact, in a set of 17 smoking and nonsmoking
patients, a wide drug concentration range was found
[0.8 –71 ␮g/L; mean (SD), 34 (4) ␮g/L for doses between
5 and 10 mg/24 h], consistent with the large variability of
OLZ pharmacokinetics reported previously (16 ).
The high sensitivity shown by this assay makes it
suitable for measuring the extremely low concentrations
of OLZ in schizophrenic patients who are heavy smokers.
It could also potentially be helpful in clarifying the
clinically relevant role played by smoking, through increases in the clearance of OLZ via CYP1A2, and in
demonstrating the marked variability of the antipsychotic
response in schizophrenic patients (2 ). Likewise, the
method could potentially be very useful for monitoring
those patients who are given other drugs capable of
inducing OLZ metabolism (e.g., anticonvulsants and barbiturates).
This work was partially supported by Grant PI020406
from Fondo de Investigacio´ n Sanitaria, Instituto de Salud
Carlos III, Ministerio de Sanidad y Consumo, Madrid,
Spain; Grant IPR00C054 from Junta de Extremadura,
Me´ rida, Spain; Grant 02/0024 from Consejeria de Sanidad
y Consumo, Junta de Extremadura, Me´ rida, Spain; and by
Convenio de Farmacovigilancia, Consejerı´a de Sanidad y
Consumo, Junta de Extremadura, Me´ rida, and Universidad de Extremadura, Badajoz, Spain. The IS (LY170222)
and OLZ were kind gifts from Eli Lilly and Company
(Indianapolis, IN).
References
1. Green B. Focus on olanzapine. Curr Med Res Opin 1999;15:79 – 85.
2. Carrillo JA, Herraiz AG, Ramos SI, Gervasini G, Vizcaino S, Benitez J. Role of
the smoking-induced CYP1A2 and polymorphic CYP2D6 in clinical response
to olanzapine. J Clin Psychopharmacol 2003;23:119 –27.
3. Gex-Fabry M, Balant-Gorgia AE, Balant LP. Therapeutic drug monitoring of
olanzapine: the combined effect of age, gender, smoking, and comedication.
Ther Drug Monit 2003;25:46 –53.
4. Boulton DW, Markowitz JS, DeVane CL. A high-performance liquid chromatography assay with ultraviolet detection for olanzapine in human plasma
and urine. J Chromatogr B Biomed Sci Appl 2001;759:319 –23.
5. Llorca PM, Coudore F, Corpelet C, Buyens A, Hoareau M, Eschalier A.
Integration of olanzapine determinations in a HPLC-diode array detection
system for routine psychotropic drug monitoring. Clin Chem 2001;47:1719 –
21.
6. Olesen OV, Linnet K. Determination of olanzapine in serum by highperformance liquid chromatography using ultraviolet detection considering
the easy oxidability of the compound and the presence of other psychotropic
drugs. J Chromatogr B Biomed Sci Appl 1998;714:309 –15.
7. Dusci LJ, Peter Hackett L, Fellows LM, Ilett KF. Determination of olanzapine
in plasma by high-performance liquid chromatography using ultraviolet
absorbance detection. J Chromatogr B Anal Technol Biomed Life Sci
2002;773:191–7.
2091
8. Raggi MA, Casamenti G, Mandrioli R, Volterra V. A sensitive high-performance liquid chromatographic method using electrochemical detection for
the analysis of olanzapine and desmethylolanzapine in plasma of schizophrenic patients using a new solid-phase extraction procedure. J Chromatogr
B Biomed Sci Appl 2001;750:137– 46.
9. Aravagiri M, Ames D, Wirshing WC, Marder SR. Plasma level monitoring of
olanzapine in patients with schizophrenia: determination by high-performance liquid chromatography with electrochemical detection. Ther Drug
Monit 1997;19:307–13.
10. Catlow JT, Barton RD, Clemens M, Gillespie TA, Goodwin M, Swanson SP.
Analysis of olanzapine in human plasma utilizing reversed-phase highperformance liquid chromatography with electrochemical detection. J Chromatogr B Biomed Appl 1995;668:85–90.
11. Berna M, Shugert R, Mullen J. Determination of olanzapine in human plasma
and serum by liquid chromatography/tandem mass spectrometry. J Mass
Spectrom 1998;33:1003– 8.
12. Kollroser M, Schober C. Direct-injection high performance liquid chromatography ion trap mass spectrometry for the quantitative determination of
olanzapine, clozapine and N-desmethylclozapine in human plasma. Rapid
Commun Mass Spectrom 2002;16:1266 –72.
13. Berna M, Ackermann B, Ruterbories K, Glass S. Determination of olanzapine
in human blood by liquid chromatography-tandem mass spectrometry.
J Chromatogr B Biomed Sci Appl 2002;767:163– 8.
14. Kassahun K, Mattiuz E, Nyhart E Jr, Obermeyer B, Gillespie T, Murphy A, et
al. Disposition and biotransformation of the antipsychotic agent olanzapine
in humans. Drug Metab Dispos 1997;25:81–93.
15. Lohr JB, Flynn K. Smoking and schizophrenia. Schizophr Res 1992;8:93–
102.
16. Callaghan JT, Bergstrom RF, Ptak LR, Beasley CM. Olanzapine. Pharmacokinetic and pharmacodynamic profile. Clin Pharmacokinet 1999;37:177–
93.
17. Bogusz MJ, Kruger KD, Maier RD, Erkwoh R, Tuchtenhagen F. Monitoring of
olanzapine in serum by liquid chromatography-atmospheric pressure chemical ionization mass spectrometry. J Chromatogr B Biomed Sci Appl 1999;
732:257– 69.
18. Olesen OV, Linnet K. Olanzapine serum concentrations in psychiatric
patients given standard doses: the influence of comedication. Ther Drug
Monit 1999;21:87–90.
19. Annesley TM. Ion suppression in mass spectrometry. Clin Chem
2003;49:1041– 4.
DOI: 10.1373/clinchem.2003.022517
Anti-Tissue Transglutaminase Antibodies in Arthritic
Patients: A Disease-specific Finding? Antonio Picarelli,1*
Marco Di Tola,1 Luigi Sabbatella,1 Stefania Vetrano,1 Maria
Cristina Anania,1 Antonio Spadaro,2 Maria Laura Sorgi,2 and
Egisto Taccari2 (1 Gastroenterological Unit, Department of
Clinical Sciences, and 2 Rheumatological Unit, Department of Medical Therapy; University “La Sapienza”,
151-00161 Rome, Italy; * address correspondence to this
author at: Department of Clinical Sciences, Policlinico
“Umberto I”, University of Rome “La Sapienza”, Viale del
Policlinico, 155-00161 Rome, Italy; fax 390649970524, email [email protected])
Tissue transglutaminase (tTG), widely distributed in human organs, is a multifunctional enzyme involved in the
cross-linking of extracellular matrix proteins, fibrogenesis, and wound healing (1 ). Recently, tTG has been
proposed as the autoantigen of anti-endomysial antibodies (EMAs) (2 ), a serologic marker of celiac disease (CD)
(3 ). Use of anti-tTG antibodies has been advocated in the
diagnostic work-up of CD (4 ), although positive results
have been reported in patients with other intestinal disorders, such as inflammatory bowel disease (5–7 ).
The aim of the present investigation was to evaluate the
2092
Technical Briefs
occurrence of anti-tTG-positive results in a cohort of
arthritic patients, in whom the target organ is located at a
distance from the intestine. Changes in anti-tTG antibodies were also investigated in patients with rheumatoid
arthritis (RA) during treatment with methotrexate (MTX),
a drug previously shown to decrease rheumatoid factor,
soluble interleukin-2 receptor, and interleukin-6 (8 ).
In this retrospective study, we enrolled 203 patients
[121 males and 82 females; mean (SD) age, 51.4 (14.1)
years; range, 17–76 years] attending our Rheumatological
Unit in 1998 –2000 and presenting without clinical signs
and symptoms suggestive of CD. Of these patients, 183
belonged to three different groups: 74 had RA according
to the American Rheumatism Association criteria (9 ); 67
had psoriatic arthritis (PsA) according to the European
Spondyloarthropathy Study Group criteria (10 ); and 42
had ankylosing spondylitis (AS) according to modified
New York criteria (11 ). The remaining 20, having knee,
hand, or hip osteoarthritis (OA) according to the American College of Rheumatology criteria (12–14 ), were considered as disease controls. Sixty untreated CD patients
[25 males and 35 females; mean (SD) age, 42.5 (11.2) years;
range, 19 –70 years], diagnosed according to the current
criteria (15 ), were selected as disease controls. Fifty-four
blood donors [26 males and 28 females; mean (SD) age,
40.2 (12.8) years; range, 20 – 67 years] were enrolled as
healthy controls.
Serum was collected from all patients, and synovial
fluid (SF) was obtained by knee aspiration from 68
arthritic patients (33 with RA, 26 with PsA, 9 with OA) for
diagnostic proposes. All procedures followed in this
study were in accordance with the ethical standards of the
responsible institutional committee on human experimentation.
IgA anti-tTG antibodies were measured by an enzyme
immunoassay (EIA) in which microtiter plate wells were
coated with recombinant human (rh) tTG (Eurospital).
According to the manufacturer’s instructions, serum was
diluted 1:26, and SF was diluted 1:5. Absorbance was
measured at 450 nm (A450 nm). For the intraassay variation,
10 sera were tested four times in the same run; the CV
(range) was 0.9 –3.6%. For the interassay variation, 10 sera
were tested in three different runs, and the CV (range)
was 2.2–7.7%. The mean anti-tTG value of the healthy
controls ⫹ 3 SD (A450 nm ⫽ 0.2) was used as the cutoff to
identify positive results. Positive anti-tTG concentrations
were classified as high (A450 nm ⬎0.57), observed only in
CD patients, and low (A450 nm ⫽ 0.2– 0.57), observed in
both CD and arthritic patients. In 40 randomly selected
arthritic patients (10 with RA, 10 with PsA, 10 with AS,
and 10 with OA), anti-tTG antibodies were also measured
by another EIA, in which wells were coated with native
human (nh) tTG isolated from erythrocytes (INOVA Diagnostics Inc.).
IgA EMAs were identified by indirect immunofluorescence on sections of monkey esophagus (Eurospital).
According to the manufacturer’s instructions, serum was
diluted 1:5 and incubated for 30 min, whereas undiluted
SF was incubated for 1 h. The presence of EMAs, evi-
denced by reticulin-like staining of smooth-muscle bundles, was evaluated by two observers (agreement rate,
98.8%) unaware of the patients’ conditions.
Forty-five of the 74 RA patients were treated with MTX,
administered in three oral doses of 2.5 mg for a total of 7.5
mg/week. If after each month of treatment clinical improvement was not satisfactory, an additional dose of 2.5
mg was prescribed, up to a maximum dose of 15 mg/
week (8 ). After 6 months of treatment, anti-tTG antibodies
were again evaluated.
The statistical significance of differences was determined by the Mann–Whitney test for independent data
and by the Wilcoxon test for paired data. Spearman rank
correlation was used to evaluate correlations. P values
ⱕ0.05 were considered significant. All statistical evaluations were carried out using Statgraphic (STSC Inc.) and
GraphPad Prism (GraphPad Inc.) software.
We found markedly increased anti-tTG antibody concentrations in 40 of 60 CD patients (67%). Low-positive
values were detectable in 16 of 60 CD (27%), 31 of 74 RA
(42%), 23 of 67 PsA (34%), and 14 of 42 AS patients (33%).
Low-positive values were also detectable in 2 of 20 OA
patients (10%) and 1 of 54 healthy controls (2%). We found
no anti-tTG antibodies in four CD patients (6%; Fig. 1).
Anti-tTG antibodies were significantly higher in CD, RA,
PsA, AS, and OA patients (P ⬍0.001 for each) than in
healthy controls. In CD patients, anti-tTG antibodies were
also significantly higher (P ⬍0.001 for each) than in RA,
PsA, AS, and OA patients. Furthermore, in OA patients,
anti-tTG antibodies were significantly lower than in RA
(P ⫽ 0.009), PsA (P ⫽ 0.018), and AS patients (P ⫽ 0.044),
whereas we found no significant differences among RA,
Fig. 1. Serum IgA anti-tTG absorbance in patients with CD or arthritis
and in healthy controls.
The number of patients, quartile cutpoints (first and third quartile), and medians
of the anti-tTG absorbance values are given. Absorbance values ⬍0.2 were
considered negative, values between 0.2 and 0.57 were considered low positive,
and values ⬎0.57 were considered high positive.
Clinical Chemistry 49, No. 12, 2003
PsA, and AS patients. Serum EMAs were detectable in all
60 CD patients, whereas in contrast, we found no EMAs in
any of the arthritic patients or healthy controls.
Anti-tTG antibody concentrations in the sera and SF of
the 68 arthritic patients who underwent knee aspiration
were significantly correlated (r ⫽ 0.69; P ⬍0.001). No
EMAs were found in any of the SF samples examined.
After 6 months of MTX treatment, serum anti-tTG
antibodies decreased significantly in the 45 RA patients
who underwent this procedure (P ⬍0.001).
All above-mentioned anti-tTG antibody concentrations
were measured by use of microtiter plate wells coated
with rhtTG. Wells coated with nhtTG were also used for
serum samples from 40 randomly selected arthritic patients, and the results were compared. Antibody concentrations measured with rhtTG and nhtTG were significantly correlated (r ⫽ 0.917; P ⬍0.0001; Fig. 2 in the Data
Supplement that accompanies the online version of this
Technical Brief at http://www.clinchem.org/content/
vol49/issue12/).
Although anti-tTG antibodies are currently used in the
diagnostic work-up of CD, some authors have demonstrated the presence of anti-tTG-positive results in patients with inflammatory bowel disease (5–7 ), chronic
liver disease (5, 7, 16 ), insulin-dependent diabetes mellitus (5 ), and heart failure (17 ), suggesting a limited specificity. Other authors have suggested that use of rhtTG as
the EIA substrate, instead of the guinea pig tTG, could
reduce the rate of false positives in patients with chronic
liver disease and insulin-dependent diabetes mellitus
(18, 19 ). Some of the above-mentioned studies, however,
have been performed with rhtTG (7, 16, 17 ).
In the present investigation, we found increased antitTG antibodies in ⬃40% of patients with RA, PsA, and AS.
In OA patients, anti-tTG antibody concentrations were
halfway between those of arthritic patients and healthy
controls, whereas in contrast, EMAs were not found in
any of the above-mentioned patients. We also identified a
suspicious zone that included low-positive anti-tTG values from both arthritic and CD patients, suggesting that
only high concentrations are strongly associated with CD,
whereas lower titers may not be disease specific. It is
unclear whether EIA positivity corresponds to the real
presence of anti-tTG antibodies in arthritic patients. However, because anti-tTG values decreased after 6 months of
MTX treatment and serum/SF antibody concentrations
correlated between them, positive anti-tTG results appear
to be an arthritic disease-associated event.
On the other hand, increased tTG has been demonstrated in the intestines of patients with CD and Crohn
disease (6, 20 ). This antigenic overexpression could explain, at least in part, the anti-tTG induction observed in
these patients. Likewise, Rosenthal et al. (21 ) have demonstrated a relationship between tTG and pathologic
mineralization in articular chondrocytes, which in turn
leads to the development of destructive arthritis. Thus a
mechanism similar to those in the intestine could occur in
synovial tissue.
If anti-tTG antibodies are present in EMA-negative
2093
arthritic patients, it is possible that tTG is not the only
antigen of the EMAs. Lock et al. (22 ) have demonstrated
that although tTG immunoabsorption is enough to abrogate anti-tTG EIA reactivity, it failed to completely abolish
EMA binding activity. Another study showed the existence of EMA antigenic epitopes not related to tTG (23 ).
Furthermore, using a proteomic approach, Stulik et al.
(24 ) recently identified new CD autoantigens, such as
ATP-synthase ␤-chain and enolase-␣, whereas in contrast,
other authors have demonstrated that EMAs are unable to
bind tTG-knockout tissues (25 ). Although the rhtTG and
nhtTG used in the present study are species specific,
EIA-positive results could also subtend antibodies directed against impurities of the substrate used for coating
plate wells. To verify whether EIA-positive results correspond to the real presence of anti-tTG antibodies in
arthritic patients, other molecular investigations are necessary.
In conclusion, our data demonstrate that both the
serum and SF of arthritic patients can give anti-tTGpositive results. This feature should be considered before
anti-tTG antibody tests are used; furthermore, EMAs
should always be evaluated when CD is suspected, especially in patients with known arthritis.
This study was partially supported by the nonprofit
society CELIDIA 2001 and by the IRCCS Ospedale Maggiore (Milan, Italy), research project “Celiac disease: develop of guidelines for the screening, diagnosis and study
of the pathogenetic mechanisms”. We are grateful to
Marian Shields for help with the English style.
References
1. Greenberg CS, Birckbichler PJ, Rice RH. Transglutaminases: multifunctional
cross-linking enzymes that stabilize tissues. FASEB J 1991;5:3071–7.
2. Dieterich W, Ehnis T, Bauer M, Donner P, Volta U, Riecken EO, et al.
Identification of tissue transglutaminase as the auto-antigen of celiac
disease. Nat Med 1997;7:797– 801.
3. Fasano A, Catassi C. Current approaches to diagnosis and treatment of
celiac disease: an evolving spectrum. Gastroenterology 2001;120:636 –51.
4. Sulkanen S, Halttunen T, Laurila K, Kolho KL, Korponay-Szabo IR, Sarnesto
A, et al. Tissue transglutaminase autoantibody enzyme-linked immunosorbent assay in detecting celiac disease. Gastroenterology 1998;115:
1322– 8.
5. Koop I, Ilchmann R, Izzi L, Adragna A, Koop H, Barthelmes H. Detection of
autoantibodies against tissue transglutaminase in patients with celiac
disease and dermatitis herpetiformis. Am J Gastroenterol 2000;95:2009 –
14.
6. Farrace MG, Picarelli A, Di Tola M, Sabbatella L, Marchione OP, Ippolito G, et
al. Presence of anti-tissue transglutaminase antibodies in inflammatory
intestinal diseases: an apoptosis-associated event? Cell Death Differ 2001;
8:767–70.
7. Carroccio A, Vitale G, Di Prima L, Chifari N, Napoli S, La Russa C, et al.
Comparison of anti-transglutaminase ELISAs and an anti-endomysial antibody assay in the diagnosis of celiac disease: a prospective study. Clin
Chem 2002;48:1546 –50.
8. Spadaro A, Taccari E, Riccieri V, Sensi F, Sili Scavalli A, Zoppini A.
Relationship of soluble interleukin-2-receptor and interleukin-6 with classspecific rheumatoid factors during low dose methotrexate treatment in
rheumatoid arthritis. Rev Rhum (Engl Ed) 1997;64:89 –94.
9. Arnett FR, Edworthy SM, Bloch DA. The American Rheumatism Association
1987 revised criteria for classification of rheumatoid arthritis. Arthritis
Rheum 1988;31:315–24.
10. Dougados M, van der Linden S, Juhlin R, Huitfeldt B, Amor B, Calin A, et al.
The European Spondyloarthropathy Study Group preliminary criteria for the
classification of spondyloarthropathy. Arthritis Rheum 1991;34:1218 –27.
11. Van der Linden S, Valkenburg HA, Cats A. Evaluation of diagnostic criteria for
2094
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
Technical Briefs
ankylosing spondylitis. A proposal for modification of the New York criteria.
Arthritis Rheum 1984;27:361– 8.
Altman R, Asch E, Bloch D, Bole G, Borenstein D, Brandt K, et al. The
American College of Rheumatology criteria for the classification and reporting of osteoarthritis of the knee. Arthritis Rheum 1986;29:1039 – 49.
Altman R, Alarcon G, Appelrouth D, Bloch D, Borenstein D, Brandt K, et al.
The American College of Rheumatology criteria for the classification and
reporting of osteoarthritis of the hand. Arthritis Rheum 1990;33:1601–10.
Altman R, Alarcon G, Appelrouth D, Bloch D, Borenstein D, Brandt K, et al.
The American College of Rheumatology criteria for the classification and
reporting of osteoarthritis of the hip. Arthritis Rheum 1991;34:505–14.
Walker-Smith JA, Guandalini S, Schmitz J, Shmerling DH, Visakorpi JK.
Revised criteria for diagnosis of celiac disease. Arch Dis Child 1990;65:
909 –11.
Vecchi M, Folli C, Donato MF, Formenti S, Arosio E, De Franchis R. High rate
of positive anti-tissue transglutaminase antibodies in chronic liver disease.
Role of liver decompensation and of the antigen source. Scand J Gastroenterol 2003;38:50 – 4.
Peracchi M, Trovato C, Longhi M, Gasparin M, Conte D, Tarantino C, et al.
Tissue transglutaminase antibodies in patients with end-stage heart failure.
Am J Gastroenterol 2002;97:2850 – 4.
Carroccio A, Giannitrapani L, Soresi M, Not T, Iacono G, Di Rosa C, et al.
Guinea pig transglutaminase immunolinked assay does not predict celiac
disease in patients with chronic liver disease. Gut 2001;49:506 –11.
Leon F, Camarero C, R-Pena R, Eiras P, Sanchez L, Baragano M, et al.
Anti-transglutaminase IgA ELISA: clinical potential and drawbacks in celiac
disease diagnosis. Scand J Gastroenterol 2001;36:849 –53.
D’Argenio G, Biancone L, Cosenza V, Della Valle N, D’Armiento FP, Boirivant
M, et al. Transglutaminases in Crohn’s disease. Gut 1995;37:690 –5.
Rosenthal AK, Derfus BA, Henry LA. Transglutaminase activity in aging
articular chondrocytes and articular cartilage vesicles. Arthritis Rheum
1997;40:966 –70.
Lock J, Gilmour JEM, Unsworth DJ. Anti-tissue transglutaminase, antiendomysium and anti-R1-reticulin autoantibodies, the antibody trinity of
coeliac disease. Clin Exp Immunol 1999;116:258 – 62.
Uhlig HH, Lichtenfeld J, Osman AA, Richter T, Mothes T. Evidence for
existence of celiac disease autoantigens apart from tissue transglutaminase. Eur J Gastroenterol Hepatol 2000;12:1017–20.
Stulik J, Hernychova L, Porkertova S, Pozler O, Tuckova L, Sanchez D, et al.
Identification of new celiac disease autoantigens using proteomic analysis.
Proteomics 2003;3:951– 6.
Korponay-Szabo IR, Laurila K, Szondy Z, Halttunen T, Szalai Z, Dahlbom I, et
al. Missing endomysial and reticulin binding of coeliac antibodies in
transglutaminase 2 knockout tissues. Gut 2003;52:199 –204.
DOI: 10.1373/clinchem.2003.023234
Reliability of IFCC Method for Lactate Dehydrogenase
Measurement in Lithium-Heparin Plasma Samples, Ileana Herzum, Robert Bu¨ nder, Harald Renz, and Hans Gu¨ nther
Wahl* (Klinikum der Philipps–Universita¨ t Marburg, Department of Clinical Chemistry and Molecular Diagnostics,
35033 Marburg, Germany; * author for correspondence: fax
49-6421-2865594, e-mail [email protected])
In a recent Technical Brief on the IFCC-recommended
method for lactate dehydrogenase (LD) measurement,
Bakker et al. (1 ) questioned the reliability of this method
for heparin samples. According to their findings, the IFCC
method produced an excessive frequency of significantly
discordant duplicate measurements (17.8%; 27 of 152
samples) compared with the formerly applied French
Society (SFBC)-recommended method (1.4%; 2 of 140
samples). Neither increased centrifugation time and
speed or the use of plasma separators to reduce the
plasma contamination with blood cells, nor LD measurements without concurrent measurements of other analytes to eliminate possible interferences lowered the re-
ported high rate of duplicate errors. The problem was also
not attributable to the clinical chemistry analyzer because
measurements on both Hitachi 911 and 717 analyzers
(Roche Diagnostics GmbH) showed results similar to
those obtained with the previously used Modular analyzer (Roche Diagnostics GmbH). Only when serum samples or secondary tubes for centrifuged plasma samples
were used was there a significantly lower frequency of
duplicate errors: 2.6% (3 of 114) for serum and 1.1% (1 of
94) for secondary plasma tubes. The authors pointed out
that the differences in buffer composition of the IFCC (pH
9.40 ⫾ 0.05) (2 ) and SFBC (pH 7.4 –7.8) methods influenced the integrity of platelets and erythrocytes. In this
case, the presence of cells might play a role in the high
frequency of duplicate errors.
We have been measuring LD activity on three Roche
Hitachi 917 analyzers according to the recommendations
of the IFCC (2 ) in our laboratory since February 2003. The
within-run imprecisions (CV) for a heparin-plasma sample (n ⫽ 20) were 0.8% (197.4 U/L) and 0.7% (321.3 U/L),
and the between-run CV were 1.4% for the Roche Precipath U (178.3 U/L; n ⫽ 28) and 1.1% for the Roche
Precinorm U calibrators (241.0 U/L; n ⫽ 28). During this
period of time we observed no obviously erratic LD
measurements for controls or patients when several samples from the same patient where analyzed within a few
hours for follow-up. All measurements were routinely
performed with plasma samples from plastic lithiumheparin-gel tubes (Monovette prod. no. 03.1631.001;
Sarstedt). Blood sample collection was done without
vacuum drawing. After centrifugation for 10 min at 3000g,
the primary tubes were used for analysis. To reevaluate
the reliability of the IFCC-recommended LD method in
our laboratory with special regard to the reported high
frequency of duplicate errors, we performed 140 LD
duplicate measurements and studied the possible interferences from platelet contamination, cuvette and reagent
probe/stirrer carryover effects, and different primary
plasma tubes. To avoid long time delays between duplicate measurements, the analyses of the 140 samples were
performed in the same run in groups of 5 with reruns.
The statistical analysis of the results from the duplicate
measurements was calculated according to the Bland–
Altman procedure (3–5 ). Fig. 1 shows the Bland–Altman
plot with the means of the duplicate LD measurements
(U/L) plotted against the absolute differences between
duplicate measurements (U/L). For the differences the
values of the second measurements were subtracted from
the values of the first. The limits of agreement [mean (2
SD), 0.3 (26.6) U/L] (3–5 ) are shown as solid lines in Fig.
1, and the mean of the differences as a dashed line. The
frequency of duplicate errors [differences exceeding mean
(2 SD)] under these conditions was 3.6% (5 of 140). This is
more than the reported 1.4% for the SFBC method but less
than the 17.8% reported for the IFCC method (1 ). The 95%
confidence interval (⫺1.95 to 2.54 U/L; mean ⫾ 2 SE)
includes 0, so there is no evidence of systematic bias (3–5 ).
[When comparing our data with the results published by