1. health and fitness
2. disease

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Technical Briefs
evaluated in our small study sample is also a potential
limitation.
This study was supported in part by grants from Fu Wai
Hospital, the Chinese Academy of Medical Science (190),
and the People’s Republic of China (1998679) to Dr. Li.
References
1. Maseri A. Inflammation, atherosclerosis, and ischemic events: exploring the
hidden side of the moon. N Engl J Med 1997;336:1014 – 6.
2. Ross R. Atherosclerosis—an inflammatory disease. N Engl J Med 1993;
340:115–26.
3. Libby P, Ridker PM. Novel inflammatory markers of coronary risk: theory
versus practice. Circulation 1999;100:1148 –50.
4. Li J-J, Fang C-H. C-Reactive protein is not only a marker but also direct cause
of cardiovascular disease. Med Hypotheses 2004;62:499 –506.
5. Liuzzo G, Biasucci LM, Gallimore JR, Grillo RL, Rebuzzi AG, Pepys MB, et al.
The prognostic value of C-reactive protein and serum amyloid a protein in
severe unstable angina. N Engl J Med 1994;331:417–24.
6. Morrow DA, Rifai N, Antman EM. C-Reactive protein is a potent predicator of
mortality independently of and in combination with troponin T in acute
coronary syndromes: a TIMI 11A substudy. J Am Coll Cardiol 1998;31:
1460 –5.
7. Li J-J, Wang H-R, Huang C-X, Xue J-L, Li G-S. Enhanced response of blood
monocytes to C-reactive protein in patients with unstable angina. Clin Chim
Acta 2005;352:127–33.
8. Smith DA, Irving SD, Sheldon J, Kaski JC. Serum levels of the antiinflammatory cytokine interleukin-10 are decreased in patients with unstable
angina. Circulation 2001;104:746 –50.
9. Alam SE, Nasser SS, Fernainy KE, Habib AA, Badr KF. Cytokine imbalance in
acute coronary syndrome. Curr Opin Pharmacol 2004;4:166 –70.
10. Heeschen C, Dimmeler S, Hamm CW, Fichtlscherer S, Boersma E, Simoons
ML. Serum level of the anti-inflammatory cytokine interleukin-10 is an
important prognostic determinant in patients with acute coronary syndromes. Circulation 2003;107:2109 –14.
11. Mizia-Stec K, Gasior Z, Zahorska-Markiewiez B, Janowska J, Szulc A,
Jastrzebska-Maj E, et al. Serum tumor necrosis factor-␣, interleukin-2 and
interleukin-10 activation in stable angina and acute coronary syndromes.
Coron Artery Dis 2003;14:431– 8.
12. Fichtlscherer S, Breuer S, Heeschen C, Dimmeler S, Zeiher A. Interleukin-10
serum levels and systemic endothelial vasoreactivity in patients with
coronary artery disease. J Am Coll Cardiol 2004;44:50 –2.
13. Schieffer B, Bunte C, Witte J, Hoeper K, Boger RH, Schwedhelm E, et al.
Comparative effects of anti-antagonism and angiotensin converting enzyme
inhibition on markers of inflammation, platelet aggregation in patients with
coronary disease. J Am Coll Cardiol 2004;44:362– 8.
14. Plenge JK, Hernandez TL, Weil KM, Poirier P, Grunwald GK, Marcovina SM,
et al. Simvastatin lowers C-reactive protein within 14 days: an effect
independent of low-density lipoprotein cholesterol reduction. Circulation
2002;106:1447–52.
15. Li J-J, Chen M-Z, Chen-X, Fang C-H. Rapid effects of simvastatin on lipid
profile and C-reactive protein in patients with hypercholesterolemia. Clin
Cardiol 2003;26:472– 6.
16. Li J-J, Chen X-J. Simvastatin inhibits interleukin-6 release in human monocytes stimulated by C-reactive protein and lipopolysaccharide. Coron Artery
Dis 2003;14:329 –34.
17. Ridker PM, Rifai N, Lowenthal SP. Rapid reduction in C-reactive with
cerivastatin among 785 patients with primary hypercholesterolemia. Circulation 2001;103:1191–3.
18. Li J-J, Jiang H, Huang C-X, Fang C-H, Tang Q-Z, Xiao H, et al. Elevated levels
of plasma C-reactive protein in patients with unstable angina: its relations
with coronary stenosis and lipid profile. Angiology 2002;53:265–72.
19. Li J-J, Fang C-H, Cheng M-Z, Cheng X. Activation of nuclear factor-␬B and
correlation with elevated plasma C-reactive protein in patients with unstable
angina. Heart Lung Circ 2004;13:173– 8.
20. Davis MJ. Stability and unstability: the two faces of coronary atherosclerosis. Circulation 1996;94:2013–20.
21. Moreno PR, Falk E, Palacios IF, Newell JB, Fuster V, Fallon JT. Macrophage
infiltration in acute coronary syndromes: implications for plaque rupture.
Circulation 1994;90:775– 80.
22. Wang P. Wu P, Siegel MI, Egan RW Billa MM. Interleukin (IL)-10 inhibits
nuclear factor ␬B (NF ␬B) activation in human monocytes: IL-10 and IL-4
suppress cytokine synthesis by different mechanisms. J Biol Chem 1995;
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23. Lacraz S, Nicod LP, Chicheortiche R, Welgus HG, Dayer JM. IL-10 inhibits
metalloproteinase and stimulates TIMP-1 production in human mononuclear
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monocyte tissue factor expression in whole blood. Br J Haematol 1998;
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26. Pinderski LJ, Fishbein MP, Subbanagounder G, Fishbein MC, Kubo N,
Cheroutre H, et al. Overexpression of interleukin-10 by activated T lymphocytes inhibits atherosclerosis in LDL receptor-deficient mice by altering
lymphocyte and macrophage phenotypes. Circ Res 2002;90:1064 –71.
27. Treasure CB, Klein JL, Weintraub WS, Talley JD, Stillabower ME, Kosinski
AS, et al. Beneficial effects of cholesterol-lowering therapy on the coronary
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1995;332:481–7.
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DOI: 10.1373/clinchem.2005.049700
Pediatric Reference Intervals for Seven Common Coagulation Assays, Michele M. Flanders,1 Ronda A. Crist,2
William L. Roberts,1,3 and George M. Rodgers1,3,4* (1 ARUP
Institute for Clinical and Experimental Pathology, Salt
Lake City, UT; 2 ARUP Laboratories, Salt Lake City, UT;
3
Department of Pathology, University of Utah Health
Sciences Center, Salt Lake City, UT; 4 Department of
Medicine, University of Utah Health Sciences Center, Salt
Lake City, UT; * address correspondence to this author at:
Division of Hematology, University of Utah Health Sciences Center, 50 North Medical Dr., Salt Lake City, UT
84132; fax 801-585-5469, e-mail [email protected].
edu)
Accurate interpretation of pediatric coagulation tests is
complicated by the fact that reference intervals for many
assays differ from those for adults. In 1992, Andrew et al.
(1 ) reported childhood coagulation reference intervals for
ages 1–5, 6 –10, and 11–16 years with a minimum of 4 and
maximum of 7 individuals at each age, with 20 –50 per age
group. These results have served as the basis for most
pediatric coagulation reference intervals for over a decade. However, with the development of newer reagents,
methodologies, and instruments to measure coagulation
analytes, results from this study, which was limited by its
small size, may be less relevant.
In 2002, we initiated a project to collect blood and urine
samples from healthy children 7–17 years of age, with the
goal of establishing pediatric reference intervals for many
laboratory tests (2 ). The purpose of this study was to
determine pediatric reference intervals for 7 coagulation
tests associated with common bleeding disorders.
Samples were drawn for reference interval studies from
902 healthy children, ages 7–17 years. All were healthy,
had no history of bleeding or thrombotic disorders, and
were taking no medications for at least 2 weeks before
specimen collection. Informed consent was obtained from
parents, and the study was approved by the University of
Utah Institutional Review Board. Samples used to establish adult reference intervals were purchased from George
1739
Clinical Chemistry 51, No. 9, 2005
King Bio-Medical or Precision Biologic, or were collected
from local volunteers.
Blood was obtained by clean venipuncture; a pilot tube
was drawn first. An exact ratio of 9 volumes of blood to 1
volume of anticoagulant (32 g/L citrate) was maintained.
Specimens were centrifuged immediately at 3000g for 20
min at room temperature, aliquoted, and frozen at
⫺80 °C.
Prothrombin time (PT); partial thromboplastin time
(PTT); factors VIII, IX, and XI; and von Willebrand factor
(vWF) antigen were assayed on the STA-R coagulation
analyzer (Diagnostica Stago). Ristocetin cofactor activity
(RCF) assays were performed on the BCS (Dade Behring).
All coagulation assays were done according to manufacturer specifications and laboratory standards. Coagulation screening tests included in the study were the PT
(Neoplastin Cl⫹; Diagnostica Stago) and PTT (STA-PTTa;
Diagnostica Stago) assays, and results were measured in
seconds with a clot-based methodology. Intrinsic factors
VIII, IX, and XI [STA-PTTa (Diagnostica Stago) and factordeficient plasma (HRF Inc.)] were measured by a modified activated PTT (APTT) (3 ). A calibration curve was
generated by use of dilutions of a commercial calibrator
and Owrens buffer, and the results were calculated from
this curve. RCF (BC vonWillebrand reagent; Dade Behring) was assayed by use of stabilized platelets added to a
sample containing vWF (ristocetin cofactor), which causes
platelet agglutination and decreased absorbance in the
presence of ristocetin (4 ). The vWF antigen (STA LIATEST vWF; Diagnostica Stago) was measured by microlatex particle-mediated immunoassay (5 ). In this assay, a
suspension of latex microparticles covalently bound to a
monoclonal antibody specific for vWF was mixed with
the test plasma. Agglutination of the latex microparticles
induces an increase in turbidity, producing an increase in
absorbance that is proportional to the vWF antigen concentration present in the sample.
For each assay described, calibration and control sam-
Table 1. Summary of pediatric reference intervals.a
Age, years
n
PT, s
Lower limit
Upper limit
Median
PTT, s
Lower limit
Upper limit
Median
Factor VIII, %
Lower limit
Upper limit
Median
Factor IX, %
Lower limit
Upper limit
Median
Factor XI, %
Lower limit
Upper limit
Median
RCF, %
Lower limit
Upper limit
Median
Mean
vWF antigen, %
Lower limit
Upper limit
Median
Mean
a
16–17
Adultb
150
120
7–9
10–11
12–13
14–15
245
164
164
164
13.0 (12.6–13.1)
15.4c (15.2–15.7)
14
13.0 (12.7–13.1)
15.6c (15.2–16.2)
14
13.0 (12.8–13.2)
15.2c (14.8–15.7)
14
12.8 (12.4–13.0)
15.4c (15.2–15.8)
14
12.6 (12.3–12.4)
15.7c (15.2–16.2)
13.9
27 (26–28)
38 (37–40)
31
27 (26–28)
38 (37–40)
31
27 (26–27)
39 (36–40)
31
26 (25–27)
36 (35–44)
31
26 (24–27)
35 (35–38)
31
76 (70–80)
199c (187–219)
122
80 (70–89)
209c (186–268)
133
72 (63–87)
198c (182–218)
131
69 (56–81)
237c (208–311)
123
63 (57–73)
221c (195–239)
122
56 (45–63)
191 (175–222)
106
70 (68–72)
133c (129–150)
91
72 (60–76)
149c (136–164)
102
73 (70–78)
152c (144–187)
103
80 (77–83)
161 (152–183)
110
86 (74–88)
176 (167–194)
118
78 (58–85)
184 (165–210)
119
70 (65–73)
138 (131–154)
99
66 (61–74)
137 (131–154)
101
68 (53–70)
138c (126–148)
91
57 (51–65)
129c (119–161)
91
65 (55–70)
159 (136–177)
92
56 (54–68)
153 (144–165)
102
52 (43–56)
176 (154–223)
88
94
60 (43–65)
195c (178–258)
101
108
50 (36–58)
184c (173–232)
101
105
50 (39–58)
203c (185–260)
103
107
49 (44–56)
204 (197–252)
97
101
51 (49–58)
215 (132–243)
75
87
62 (50–65)
180c (164–197)
101
105
63 (61–69)
189c (170–244)
109
116
60 (47–70)
189c (179–200)
108
114
57 (49–61)
199c (189–238)
108
113
50 (42–58)
205 (167–252)
104
108
52 (46–60)
214 (159–308)
94
100
12.3 (11.9–12.2)
14.4 (14.3–14.6)
13.2
26 (25–26)
38 (36–38)
31
The 90% confidence interval for each reference limit is given in parentheses.
Adult reference intervals for RCF and vWF antigen were calculated from samples drawn from Utah donors only as described in the text (n ⫽ 78).
c
The Z-test of the mean, the ratio test for the SD, or both are statistically different from the adult reference intervals.
b
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Technical Briefs
Fig. 1. Data from pediatric populations for 7 coagulation assays.
Shaded boxes indicate the pediatric reference intervals (central 95% interval) for each age group. Within the box is a solid line that represents the median of the group.
Dashed lines represent the median and central 95% interval of the adult reference data compared with the pediatric intervals. In panel B, the median is the same for
children ages 7–15 years and adults and is indicated by a dashed line only.
Clinical Chemistry 51, No. 9, 2005
ples were assayed daily. Testing was performed in
batches of 20 specimens. For each batch, a total of 20 (10
male/10 female) randomly chosen samples were thawed
for 7 min in a 37 °C waterbath and run concurrently. Data
from the 902 samples were grouped by age: 7–9, 10 –11,
12–13, 14 –15, and 16 –17 years. Data were grouped with a
minimum of 164 individuals in each age group, allowing
the establishment of reference intervals by a nonparametric method (NCCLS C28-A) (6 ). Mean (SD) values of each
age group were compared with those for adults; if the
Z-test of the means (calculated Z is greater than critical Z),
or if the SD ratio (adult SD/pediatric SD) was ⬎1.5,
separate reference intervals were warranted (6 ). We used
120 normal adult samples to establish the adult reference
intervals (6 ). Of these, 78 were ARUP (Utah) donors. The
mean adult ages for ARUP donors and commercial donors were 32 and 36 years, respectively. The adult age
range for local donors was 20 –55 years, and that for
purchased plasmas was 18 –55 years. Statistical calculations were performed with EP Evaluator, release 5 (David
G. Rhoads Associates).
Pediatric reference intervals for the central 95% (age
group 7–9, n ⫽ 245; all other age groups, n ⫽ 164) and 90%
confidence intervals are summarized in Table 1. Confidence intervals were calculated by EP Evaluator from
Table 8 of NCCLS C28-A. Individual data for each analyte
are shown in Fig. 1 (panels A–G), and both pediatric and
adult reference intervals and median values are indicated.
For RCF and vWF antigen, mean values are also included
because the median value most likely includes data from
a blood type O donor, which caused the value to be
skewed low.
The median PT for pediatric samples was 14.0 s, nearly
1 s longer than the adult mean of 13.2 s. Although the
median PT for pediatric samples decreased slightly at age
16 –17 to 13.9 s, the intervals for all pediatric age groups
were statistically significantly different from that for
adults for this analyte. However, pediatric PTT intervals
did not differ from that for adults (Table 1).
For factor VIII, RCF, and vWF antigen, all pediatric age
groups had higher median values than those of adults.
The lower limits of the reference intervals for factor VIII
and vWF antigen were markedly higher at younger ages.
However, by ages 16 –17 years, reference limits were
comparable to those for adults. vWF antigen was slightly
lower (2%). RCF lower limits were slightly higher (1%) at
age 7–9 years and significantly higher (9%) at age 10 –11
years, whereas the limits for ages 12–17 were slightly
lower than that for adults. For all pediatric age groups, the
upper limits for RCF and vWF antigen were considerably
lower than that for adults; however, the upper limits for
factor VIII were higher than for adults. The median
pediatric factor XI concentrations were similar to that for
adults, but the reference intervals had a higher lower
limit. Vitamin K-dependent coagulation factor IX concentrations were significantly different in the younger years
of life. Pediatric concentrations of factor IX trended upward until ages 16 –17, when adult values were reached.
In this study, 0.44% of the presumed “normal” samples
1741
were found to be abnormal. Three children had factor XI
deficiency (concentrations ⬍34%), and 1 had von Willebrand disease (RCF ⬍29% and vWF antigen ⬍35%). These
values were well outside established reference intervals
and were excluded when we calculated the current pediatric reference intervals.
Most coagulation laboratories rely on the published
pediatric reference intervals reported by Andrew et al. (1 )
over a decade ago. That study reported results for a
maximum of 50 children per age group, and each age
group spanned 5 years (1–5, 6 –10, and 11–16 years). Their
study did not identify significant differences between
children of any age and the adult population for the PT,
PTT, factor VIII, and vWF antigen assays. Factor IX
concentrations did show age dependence, as did factor XI
in the older pediatric age group (11–16 years). RCF was
not reported in the study by Andrew et al. (1 ).
In our study, we found age-dependent differences
between children and adults for the PT assay; factors VIII,
IX, and XI; RCF; and vWF antigen (Table 1). The major
reason we detected these differences and the earlier study
did not is probably the larger number of individuals and
the more restrictive age groups in our study.
Our results have implications for the interpretation of
pediatric coagulation assays. For example, a PT that is
prolonged by 1 s in a child, based on an adult reference
interval, is probably physiologic and does not indicate a
disease state. Similarly, our new findings for the vWF
panel component reference intervals (factor VIII, RCF,
and vWF antigen) could lead to more accurate identification of von Willebrand disease in childhood.
One genetic modifier of plasma vWF concentrations is
the ABO blood group; individuals with type O have
⬃25% lower vWF concentrations than do those with a
non-O blood type (7 ). Although our pediatric samples
were not analyzed for ABO blood type, blood bank data
from Utah blood donors indicates that a similar but
slightly lower percentage have blood type O compared
with the overall US blood donor pool (35% vs 45%).
Therefore, to calculate the vWF and the RCF adult reference intervals, we omitted out-of-state donor plasma
samples and relied on Utah adult donors to match Utah
pediatric donors. Our adult reference interval values for
vWF and RCF are slightly higher than the corresponding
pediatric intervals, a finding similar to that reported by
Andrew et al. (1 ).
In conclusion, we found several significant differences
between pediatric and adult coagulation reference intervals, supporting the need to establish new pediatric
reference intervals. Our study has established reference
intervals based on the use of newer reagents, methodologies, and instrumentation currently used in the clinical
laboratory. Future analysis of our stored plasma samples
will focus on the remaining clinically relevant coagulation
factors [fibrinogen, prothrombin (factor II), and factors V,
VII, and X] and the natural anticoagulants (protein C,
protein S, and antithrombin). These studies should improve the diagnosis of childhood bleeding and thrombotic
disorders.
1742
Technical Briefs
ARUP Institute for Clinical and Experimental Pathology
provided support for this project. Reagents and instrumentation were provided in part by Diagnostica Stago.
We would like to thank John Simmons, LaRayne Burrows,
and Ashley Widdison for their recruitment of participants
and collection of samples.
References
1. Andrew M, Vegh P, Johnston M, Bowker J, Ofosu F, Mitchell L. Maturation of
the hemostatic system during childhood. Blood 1992;80:1998 –2005.
2. Children’s Health Improvement through Laboratory Diagnostics (CHILDx).
Pediatric reference interval project. http://www.childx.org/ (accessed February 2005).
3. Sirridge MS. Laboratory evaluation of hemostasis and thrombosis. Philadelphia, PA: Lea & Febiger, 1974:130 –3.
4. Dade Behring. Ristocetin cofactor activity by BCS analyzer [Package Insert].
Deerfield, IL: Dade Behring, January 1999.
5. Diagnostica Stago. Von Willebrand factor antigen by STA analyzers [Package
Insert]. Parsippany, NJ: Diagnostica Stago, November 1997.
6. National Committee for Clinical Laboratory Standards. How to define and
determine reference intervals in the clinical laboratory; approved guideline.
NCCLS document C28-A2, Vol. 15, No. 4. Wayne, PA: NCCLS, 2000.
7. Gill JC, Endres-Brooks J, Bauer PJ, Marks WJJR, Montgomery RR. The effect
of ABO blood group on the diagnosis of von Willebrand disease. Blood
1987;69:1691–5.
DOI: 10.1373/clinchem.2005.050211
Purine Metabolites in Gout and Asymptomatic Hyperuricemia: Analysis by HPLC–Electrospray Tandem
Mass Spectrometry, Jiyuan Zhao,1 Qionglin Liang,1 Guoan
Luo,1* Yiming Wang,1 Yanjia Zuo,1 Ming Jiang,2 Guilan Yu,2
and Ting Zhang3 (1 Analysis Center, Institute of Biomedicine, Tsinghua University, Beijing, China; 2 XieHe Hospital, Beijing, China; 3 Capital Institute of Pediatrics, Beijing,
China; * address correspondence to this author at: Analysis Center, Institute of Biomedicine Tsinghua University,
Beijing 100084, People’s Republic of China; fax 86-1062781688, e-mail [email protected])
Hyperuricemia, a serum urate concentration ⬎0.45
mmol/L (7.0 mg/dL) in men and 0.36 mmol/L (6.0
mg/dL) in women, is the biochemical hallmark of gout
(1–3 ), but many individuals with life-long hyperuricemia
do not develop gouty arthritis (4 ). Conversely, serum
urate concentrations may be within reference values in
some patients with acute gout, particularly during the
early phases of the disorder. We hypothesized that gout
patients may have a different profile of purine precursors
than do asymptomatic people with hyperuricemia and
that concentrations of these compounds may reflect disorders of purine metabolism. To our knowledge, purine
metabolites have not been studied and compared in gout
and asymptomatic hyperuricemia.
With established methods for measuring purine compounds in blood and urine (5–9 ), retention time is not
always adequate for identification of every peak because
urine and blood usually contain many interfering compounds. Quantification by tandem mass spectrometry
(MS/MS) (10, 11 ) may require an internal standard,
which often is hard to obtain.
In this study of 10 purine compounds in the blood of
gout patients and hyperuricemia patients with no gout
symptoms, we used HPLC for serum sample separation,
MS/MS for peak identification, and ultraviolet (UV) detection for quantification.
Purines were purchased from Fluka and Sigma. The
HPLC-MS experiment was performed on an Agilent 1100
series liquid chromatograph with a mass spectrometric
detector trap. Separation was carried out on a SupelcosilTM LC-18-DB column [25 cm ⫻ 4.6 mm (i.d.); 5-␮m
film thickness]. The column temperature was maintained
at 25 °C. The mobile phases were as follows: 10 mmol/L
ammonium acetate, adjusted to pH 6.5 with glacial acetic
acid (eluant A), and methanol (eluant B). The elution
gradient was as follows (flow rate, 1 mL/min): 0 to 10
min, 100% A; 10 to 16 min, 100% A to 92% A; 16 to 30 min,
92% A to 80% A; 30 to 45 min, 80% A to 0% A; 45 to 50
min, 0% A to 100% A; 50 to 60 min, equilibration with
100% A. All gradient steps were linear, and the total
analysis time, including equilibration, was 60 min. The
column eluate was monitored at 254 nm. A splitter was
used between the HPLC column and the mass spectrometer, and 100 –200 ␮L/min of eluate was introduced into
the mass spectrometer. Negative electrospray ionization
mode was used. Nitrogen was used as both nebulizing
and collision gas, and capillary voltage was maintained at
3.5 kV. Other MS conditions were as follows: nebulizer, 25.0
psi; dry gas, 8.0 L/min; dry temperature, 325 °C; target
mass, 200 m/z; compound stability, 60%; trap drive level,
50%; ICC target, 20 000. Auto-MS/MS was performed on
molecular ions of all peaks to obtain their fragment information. Compound peaks were identified by their retention
times and auto-MS/MS. Identification results for the molecular ions and the most abundant ions of 10 compounds are
listed in Table 1 of the Data Supplement that accompanies
the online version of this Technical Brief at http://www.
clinchem.org/content/vol51/issue9/. Quantification was
based on UV detection at 254 nm.
Blood samples were obtained from 49 untreated gout
patients, 12 treated gout patients, 20 patients with untreated asymptomatic hyperuricemia, 13 patients with
treated asymptomatic hyperuricemia, and 35 healthy controls. All patients and controls were males with no renal
dysfunction. All treated patients were receiving 0.1 g/day
allopurinol. All blood samples were centrifuged to obtain
serum in the hospital and sent to our laboratory, where they
were analyzed within 2 h or stored at ⫺80 °C. Before
analysis, 100 ␮L of serum was mixed with 500 ␮L of
methanol for deproteinization, centrifuged at 14 800g for 10
min, dried with gentle nitrogen at 50 °C, and then mixed
with 100 ␮L of water for HPLC-UV-MS/MS analysis.
The limits of detection and the calibration equations for
10 compounds are listed in Table 2 of the online Data
Supplement. We determined the intraassay variation of
the method (see Table 3 of the online Data Supplement)
by measuring a serum sample 7 times and a serum sample
enriched with synthetic compounds at low (100 ␮g/L),