Download at 4°C increased to 21% and 20%, respectively

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Technical Briefs
at 4 °C increased to 21% and 20%, respectively, the
differences did not reach statistical significance (P ⬎0.05).
There was also no significant difference in the 356/105
ratios for samples subjected to other preanalytical conditions.
Because we have shown that repeated freezing and
thawing of plasma samples would affect the integrity of
plasma DNA, it is logical to investigate whether freezing
and thawing of extracted DNA would also lead to fragmentation of DNA. The 201/105 and 356/105 ratios for
plasma DNA subjected to one and three cycles of additional freezing and thawing of extracted DNA are shown
in Fig. 1D. There was no significant change in DNA
concentration when the extracted DNA was frozen and
thawed up to three times.
In this study, we have shown that clotting and delayed
separation of plasma from blood cells for 24 h significantly increases the concentration and observed size of
cell-free DNA in blood samples. Moreover, we have also
shown that repeated freezing and thawing of plasma
samples, but not extracted DNA, leads to fragmentation
of DNA. Therefore, blood samples collected for investigation of the integrity of circulating DNA should be handled
within 6 h after collection. The harvested plasma should
be aliquoted into smaller portions to avoid repeated
freezing and thawing of samples. Alternatively, DNA can
be extracted from the plasma samples for storage because
DNA appears to be more resistant to fragmentation when
stored in DNA extraction solution than in plasma.
This project is supported by a Central Allocation Grant
(CUHK1/03C) from the Hong Kong Research Grants
Council.
References
1. Leon SA, Shapiro B, Sklaroff DM, Yaros MJ. Free DNA in the serum of cancer
patients and the effect of therapy. Cancer Res 1977;37:646 –50.
2. Lo YMD, Lau TK, Zhang J, Leung TN, Chang AM, Hjelm NM, et al. Increased
fetal DNA concentrations in the plasma of pregnant women carrying fetuses
with trisomy 21. Clin Chem 1999;45:1747–51.
3. Lo YMD, Leung TN, Tein MS, Sargent IL, Zhang J, Lau TK, et al. Quantitative
abnormalities of fetal DNA in maternal serum in preeclampsia. Clin Chem
1999;45:184 – 8.
4. Lo YMD, Rainer TH, Chan LY, Hjelm NM, Cocks RA. Plasma DNA as a
prognostic marker in trauma patients. Clin Chem 2000;46:319 –23.
5. Leon SA, Ehrlich GE, Shapiro B, Labbate VA. Free DNA in the serum of
rheumatoid arthritis patients. J Rheumatol 1977;4:139 – 43.
6. Chan KCA, Zhang J, Hui ABY, Wong N, Lau TK, Leung TN, et al. Size
distributions of maternal and fetal DNA in maternal plasma. Clin Chem
2004;50:88 –92.
7. Li Y, Zimmermann B, Rusterholz C, Kang A, Holzgreve W, Hahn S. Size
separation of circulatory DNA in maternal plasma permits ready detection of
fetal DNA polymorphisms. Clin Chem 2004;50:1002–11.
8. Wang BG, Huang HY, Chen YC, Bristow RE, Kassauei K, Cheng CC, et al.
Increased plasma DNA integrity in cancer patients. Cancer Res 2003;63:
3966 – 8.
9. Chan KCA, Zhang J, Chan AT, Lei KI, Leung SF, Chan LY, et al. Molecular
characterization of circulating EBV DNA in the plasma of nasopharyngeal
carcinoma and lymphoma patients. Cancer Res 2003;63:2028 –32.
Previously published online at DOI: 10.1373/clinchem.2004.046219
Biochip for K-ras Mutation Screening in Ovarian Cancer, Gerhild Fabjani,1,2 Gernot Kriegshaeuser,3 Andreas Schuetz,4
Lothar Prix,4 and Robert Zeillinger1,3* (1 Department of Obstetrics and Gynecology, Division of Gynecology, Medical
University of Vienna, Vienna, Austria; 2 Ludwig Boltzmann Institute for Gynecology and Gynecologic Oncology, Vienna, Austria; 3 ViennaLab Labordiagnostika
GmbH, Vienna, Austria; 4 Biofocus GmbH, Recklinghausen, Germany; * address correspondence to this author at: Department of Obstetrics and Gynecology, Division of Gynecology, Medical University of Vienna,
Vienna, Austria, Waehringer Guertel 18-20, 1090 Vienna,
Austria; fax 43-1-40400-7832, e-mail robert.zeillinger@
meduniwien.ac.at)
Ovarian carcinoma is the fifth most common female
cancer type and the most common cause of death from
gynecologic malignancies in the Western world (1 ). The
three members of the ras gene family, H-ras, K-ras, and
N-ras, are among the most common oncogenes associated
with human neoplasms (2 ). Mutations in the K-ras gene
are frequently found in malignant neoplasms: 90% of
adenocarcinomas of the pancreas; 50% of colon, 30% of
lung, and 50% of thyroid tumors; and 30% of myeloid
leukemia cases, respectively (3 ). K-ras-activating mutations occur in codons 12 and 13 and seldom in codon 61,
and lead to constitutive activation of the protein by
increasing GDP/GTP exchange or decreasing GTPase
activity of the protein, thus leading to increased cell
proliferation.
K-ras mutation frequencies seem to be highly related to
tumor histology. In general, K-ras mutations occur more
frequently in mucinous tumors, including borderline malignancies, than in nonmucinous tumors such as serous
carcinomas (4 – 8 ). K-ras mutations are more common in
borderline serous tumors than in serous carcinomas,
suggesting distinct etiologies (5, 9 –11 ).
A biochip application for detection of the 10 most
common mutations of K-ras codons 12 and 13 (12 ) combines mutant-enriched amplification with a highly specific hybridization protocol. The chip appears suitable for
the detection of K-ras mutations in human feces (12 ). An
improved biochip platform, called GeneStiX (ViennaLab
Labordiagnostika GmbH), is designed to meet the needs
of molecular diagnostic applications. Up to 400 different
DNA capture oligonucleotides can be immobilized on the
tip of a special plastic stick contained in a cylindrical tube.
This allows hybridization with low volumes in a closed
system (tube) and the use of standard laboratory equipment, such as a thermoshaker (Fig. 1).
To evaluate the compatibility of the GeneStiX system
with rapid mutation screening in tumor tissue, we analyzed ovarian tumor specimens for the presence of variations in the K-ras gene. We did not study K-ras codon 61
mutations because of their reported low frequency in
ovarian carcinomas (13–15 ).
We collected 85 ovarian tumor specimens from patients
seen at the Department of Obstetrics and Gynecology at
Clinical Chemistry 51, No. 4, 2005
Fig. 1. The GeneStiX system.
(A), detection principle of the GeneStiX. DNA oligonucleotides are immobilized on
the bottom surface (6 ⫻ 8 mm) of the plastic stick and hybridized with
biotinylated target molecules (e.g., PCR products). After staining with streptavidin– horseradish peroxidase, the stick is positioned in close proximity to a
charge-coupled device (CCD) camera chip. Chemiluminescence imaging is initialized by addition of a droplet of luminol substrate between the CCD chip and the
surface of the GeneStiX. The CCD chip is then cleaned by a water rinse and dried
in an airstream. (B), chemiluminescent image of a GeneStiX for K-ras mutation
genotyping. Columns 0, 1, and 9 show signals produced by controls (CTRL) for
the extraction procedure, PCR reaction, and hybridization process. The signal for
the K-ras mutation-specific probe represents a Gly-to-Val mutation in codon 12.
the University of Vienna School of Medicine between 1991
and 1997. The histologic types included 36 serous carcinomas, 10 mucinous carcinomas, 11 endometrioid carcinomas, 5 clear cell neoplasms, 5 undifferentiated carcinomas, 1 mixed tumor (serous undifferentiated), 6
nonepithelial tumors, and 11 tumors of borderline malignancy (5 mucinous, 5 serous, and 1 endometrioid). DNA
was isolated by use of commercially available DNA
extraction reagents (DNA Extraction System I; ViennaLab
Labordiagnostika GmbH) and was stored at 4 °C until
analyzed.
K-ras mutant-enriched PCR amplification and K-ras
GeneStiX hybridization were done according to the manufacturer’s protocols. Briefly, downstream primers were
785
biotin-labeled, and upstream primers were phosphorylated at the 5⬘ position. A 10-␮L portion of the PCR
products was digested with 1 ␮L of ␭-Exonuclease (New
England BioLabs, Inc.) for 30 min at room temperature
before dilution with the provided assay buffer, which
included a hybridization control oligonucleotide. The
exonuclease-treated PCR product was then transferred
to a GeneStiX tube containing 120 ␮L of hybridization buffer. Hybridization of the GeneStiX was performed
at 37 °C for 1 h in a conventional thermoshaker (Eppendorf AG) to ensure adequate temperature control and
constant mixing. Without additional washing steps, the
biochip was stained for 10 min with the provided streptavidin– horseradish peroxidase conjugate and chemiluminescence substrate. Sticks were then rinsed with 2 mL of
assay buffer and analyzed with the GeneStiX-Imager, a
chemiluminescence detector developed for use with the
GeneStiX system (Fig. 1). Images were automatically processed with the test-specific analysis software provided with
the imager, and a report was generated for each sample.
We identified 17 samples with K-ras mutations;
15 (88%) were positive for codon 12 mutations, and
2 (12%) were positive for codon 13 K-ras mutations. Seven
of 17 (41%) mutations found in our study were Asp12,
followed by 3 tumors containing the Val12 mutation, 3
tumors containing the Cys12 mutation, 2 tumors containing the Asp13 mutation, 1 containing the Arg12 mutation,
and 1 containing both the Ala12 and Asp12 mutations.
Mutations at codon 12 of the K-ras gene were present in
27% (3 of 11) of borderline tumors, 50% (5 of 10) of
mucinous ovarian carcinomas, 14% (5 of 36) of serous
carcinomas, and 18% (2 of 11) of endometrioid carcinomas
(Table 1). One serous and two mucinous borderline tumors contained mutations in codon 12. Gly-to-Asp
(GGT3 GAT) mutations at codon 12 of the K-ras gene
were common for two borderline tumors (one mucinous
and one serous), two mucinous ovarian carcinomas, one
clear cell neoplasm, one endometrioid tumor, and one
nonepithelial tumor, but not for the serous ovarian carcinomas. Remarkably, for the five serous ovarian carcinomas positive for K-ras mutations, the codon 12 Gly-to-Cys
(GGT3 TGT) mutation occurred exclusively in three cases.
K-ras mutation status was not correlated with either
FIGO stage or histologic type, whereas the presence of
K-ras mutations was significantly higher in well-differentiated ovarian tumors than in moderately or poorly differentiated ones (P ⫽ 0.026). This finding is consistent
with recently published data and supports the hypothesis
that well-differentiated ovarian tumors and moderately or
poorly differentiated ones develop along different pathways (16 –18 ).
All mutations found by GeneStiX hybridization were
confirmed by sequencing after mutant-enriched PCR amplification, which is in agreement with former validation
experiments (12 ). However, mutations were not detectable by sequencing without previous enrichment of mutants. For specificity and sensitivity, peptide nucleic acid
(PNA) mediates preferential amplification of mutant Kras sequences (19 ). We evaluated the sensitivity of the
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Technical Briefs
Table 1. Characteristics of 85 ovarian tumor specimens.
n
Ovarian cancers
Histologic type
Serous carcinoma
Mucinous carcinoma
Endometrioid carcinomasa
Clear cell neoplasms
Undifferentiated carcinoma
Nonepithelial tumor
Mixed
Differentiation grade
G1
G2
G3
FIGO
I
II
III
IV
Borderline tumors
Histologic type
Serous
Mucinous
Endometrioid
Total
a
Codon 12 Codon 13
Mutated, mutations, mutations,
n (%)
n
n
74 14 (19)
12
2
36
10
11
5
5
6
1
5 (14)
5 (50)
2 (18)
1 (20)
0 (0)
1 (17)
0 (0)
4
4
2
1
0
1
0
1
1
0
0
0
0
0
15
15
44
7 (47)
2 (13)
5 (11)
6
2
4
1
0
1
23
8
40
3
11
4 (17)
1 (13)
9 (23)
0 (0)
3 (27)
4
1
7
0
3
0
0
2
0
0
5
1 (20)
5
2 (40)
1
0 (0)
85 17 (20)
1
2
0
15
0
0
0
2
One tumor contained both the Ala12 and the Asp12 mutations.
amplification specific to K-ras mutants by use of a dilution
series of DNA mutant for K-ras (extracted from SW480
cells) mixed with wild-type DNA (Colo320 cells). Even in
a 1000-fold excess of wild-type DNA, GeneStiX hybridization unambiguously identified a single mutation.
Although a capture probe for the wild type is present
on the biochip, wild-type DNA is usually not detected
because of the presence of excess PNA, which is complementary to the wild-type capture probe. Actually, detection of wild-type DNA is already prohibited by the
PNA-clamped amplification procedure, which ideally
would generate only mutated amplification products.
However, in some cases, amplification of the wild-type
sequence is not completely suppressed, leading to a
visible band in a gel but lacking a signal for a mutation
after hybridization on the biochip. As evidenced by
sequencing, these amplification products consist of wildtype sequences obviously not suppressed by PNA clamping (12 ). Rarely, hybridization of the amplified wild-type
product can lead to an increase in background hybridization signal, mainly nonspecific for Gly12 and/or Asp13.
In this case, a relatively strong hybridization signal for the
wild-type capture probe compared with the probes for
mutant K-ras is generated, which is indicative of an
unexpected wild-type amplification during PNA-PCR.
Thus, the sensitivity of the introduced K-ras mutation
detection system is attributable to PNA-PCR, whereas
biochip hybridization is required to improve detection
specificity rather than enhancing sensitivity. To detect a
single mutation, several quality criteria, which have been
described by Prix et al. (12 ), must be fulfilled and are
automatically checked by the test-specific analysis software. Problems with the GeneStiX system occur if the
amount of mutated DNA is ⬎1000-fold lower than the
amount of wild-type DNA. In those instances, the reproducibility of the hybridization signal intensity is reduced
because of decreased sensitivity and specificity of the
PNA-PCR. Consequently, hybridization efficiency is decreased for the three spotted capture probes indicative of
a single K-ras mutation.
As in other studies, K-ras mutations were less common
in serous ovarian tumors (15%) than in mucinous lesions
(47%) (4, 6 – 8, 16, 17, 20 ) and more common in borderline
tumors (27%; 3 of 11) than in invasive cancers (19%; 14 of
74) (16 ). Additionally, we found that 40% of the investigated mucinous ovarian tissue specimens had a mutation
in K-ras codon 12, which was the highest detection
frequency in a subgroup of patients in this study and
significantly higher than the frequency of 12% for serous
ovarian tissue specimens. These findings are in line with
previously reported data (6, 8 ). In addition to serous
borderline tumors, one mucinous borderline tumor and
two mucinous ovarian carcinomas exhibited the same
mutation at codon 12 (Gly to Asp). Ovarian tumors of
borderline malignancy might represent a pathologic continuum between benign and invasive carcinoma. This
observation was made only for the mucinous subtype,
indicating that initiation and progression of serous carcinomas might be different. Interestingly, the K-ras status in
this study supports the view that some serous and mucinous borderline tumors might develop along the same
pathways. In that case, genes other than K-ras might be
involved in the development of different histologic subtypes of borderline tumors arising from benign tissues.
Alterations in the K-ras oncogene may be clinically important with respect to tumor etiology, early diagnosis,
and prognosis. Whether K-ras mutation analysis will also
have an impact on therapeutic intervention remains to be
seen.
We conclude that the GeneStiX system is well suited for
analyzing K-ras mutations in tumor tissue specimens. The
protocol presented here is fast, easy to perform, and
reproducible. The stick-in-a-tube principle is convenient
for daily use in routine diagnostics. The reports generated
by the software are sufficient for data analysis, and no
additional calculations are necessary. The results of this
study are consistent with those reported previously and
strengthen the thesis that some mucinous borderline
tumors may progress to mucinous ovarian carcinomas
based on the finding of the same K-ras mutations in both
ovarian tumor subtypes. For serous ovarian carcinomas,
K-ras status suggests an alternative tumorigenic pathway
that differs from that of mucinous ovarian carcinomas.
References
1. Landis SH, Murray T, Bolden S, Wingo PA. Cancer statistics, 1999. CA
Cancer J Clin 1999;49:8 –31.
Clinical Chemistry 51, No. 4, 2005
2. Aunoble B, Sanches R, Didier E, Bignon YJ. Major oncogenes and tumor
suppressor genes involved in epithelial ovarian cancer [Review]. Int J Oncol
2000;16:567–76.
3. Bos JL. ras oncogenes in human cancer: a review. Cancer Res 1989;49:
4682–9.
4. Enomoto T, Weghorst CM, Inoue M, Tanizawa O, Rice JM. K-ras activation
occurs frequently in mucinous adenocarcinomas and rarely in other common
epithelial tumors of the human ovary. Am J Pathol 1991;139:777– 85.
5. Teneriello MG, Ebina M, Linnoila RI, Henry M, Nash JD, Park RC, et al. p53
and Ki-ras gene mutations in epithelial ovarian neoplasms. Cancer Res
1993;53:3103– 8.
6. Ichikawa Y, Nishida M, Suzuki H, Yoshida S, Tsunoda H, Kubo T, et al.
Mutation of K-ras protooncogene is associated with histological subtypes in
human mucinous ovarian tumors. Cancer Res 1994;54:33–5.
7. Pieretti M, Cavalieri C, Conway PS, Gallion HH, Powell DE, Turker MS.
Genetic alterations distinguish different types of ovarian tumors. Int J Cancer
1995;64:434 – 40.
8. Fujita M, Enomoto T, Inoue M, Tanizawa O, Ozaki M, Rice JM, et al. Alteration
of the p53 tumor suppressor gene occurs independently of K-ras activation
and more frequently in serous adenocarcinomas than in other common
epithelial tumors of the human ovary. Jpn J Cancer Res 1994;85:1247–56.
9. Taylor RR, Linnoila RI, Gerardts J, Teneriello MG, Nash JD, Park RC, et al.
Abnormal expression of the retinoblastoma gene in ovarian neoplasms and
correlation to p53 and K-ras mutations. Gynecol Oncol 1995;58:307–11.
10. Haas CJ, Diebold J, Hirschmann A, Rohrbach H, Lohrs U. In serous ovarian
neoplasms the frequency of Ki-ras mutations correlates with their malignant
potential. Virchows Arch 1999;434:117–20.
11. Mok SC, Bell DA, Knapp RC, Fishbaugh PM, Welch WR, Muto MG, et al.
Mutation of K-ras protooncogene in human ovarian epithelial tumors of
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12. Prix L, Uciechowski P, Bockmann B, Giesing M, Schuetz AJ. Diagnostic
biochip array for fast and sensitive detection of K-ras mutations in stool. Clin
Chem 2002;48:428 –35.
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cancer. DNA Cell Biol 1993;12:623–7.
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Association of Ki-ras with amplified DNA sequences, detected in human
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DOI: 10.1373/clinchem.2004.041194
Measurement of Cystine in Urine by Liquid Chromatography–Tandem Mass Spectrometry, Joanne E. Wear*
and Brian G. Keevil (Department of Clinical Biochemistry,
Wythenshawe Hospital, Southmoor Road, Wythenshawe,
Manchester M23 9LT, United Kingdom; * author for correspondence: fax 44-0161-291-2927, e-mail joanne.wear@
smuht.nwest.nhs.uk)
Cystine is a neutral sulfur-containing amino acid involved
in a variety of important cellular functions, including
detoxification, metabolism, and protein synthesis. Cystine
787
is excreted through the kidneys, with the epithelial cells
of the renal proximal tubules absorbing ⬃99% of filtered
cystine through a high-affinity luminal transport system
in the proximal renal tubule, which also carries the dibasic
amino acids lysine, arginine, and ornithine. Genetic mutations in the genes encoding this transport system can
cause cystinuria, characterized by excessive amounts of
cystine, arginine, lysine, and ornithine in the urine (1 ) and
decreased absorption of cystine through the intestine (2 ).
The primary clinical manifestation of cystinuria is recurrent nephrolithiasis, which can lead to urinary tract
obstruction and renal insufficiency. Cystinuria accounts
for 1% of renal stones, with an incidence of 1:100 000
(3, 4 ). The renal stones are formed through nucleation of
crystals from a supersaturated solution. Treatment is
therefore aimed at lowering the urinary cystine concentration to below its solubility limit of 250 mg/L. The
solubility of cystine begins to decrease below pH 7;
therefore, alkalinization of the urine can be used to
prevent the recurrence of renal stones. Other treatments
used in cystinuria include low-protein diets (⬍0.8
g 䡠 kg⫺1 䡠 day⫺1) and low-sodium diets, as the presence of
sodium in the urine decreases the solubility of cystine (5 ).
Chelating agents such as captopril have also been used
(6 ).
It is important to identify patients with cystinuria to
enable the correct treatment to be given to prevent recurrent renal stone formation. Different methods have been
used for this, including spectrophotometric measurement
of cysteine in urine with phosphotungstate used as a
chromogen and by HPLC followed by ultraviolet detection. These methods are time-consuming, however: the
HPLC method requires 2 h for the analysis of one sample
(7 ), and the phosphotungstate method does not allow
accurate quantification. Often amino acid analyzers are
used for the diagnosis of cystinuria in metabolic centers.
An alternative method is needed to allow the fast, robust
quantification of urinary cystine to identify those individuals at risk of recurrent stone formation. We have chosen
reversed-phase HPLC coupled to tandem mass spectrometry (LC-MS/MS) to measure cystine in urine, as we
believe that this provides the necessary sensitivity, specificity, and speed to allow quantification of cystine in
urine for routine measurement.
We prepared a 10 mg/L stock solution of d4-deuterated
cystine (QMx Laboratories Ltd.) in 500 mmol/L NaOH.
This was diluted in distilled water to a concentration of 50
mg/L, and 30 ␮L of this was added as an internal
standard to 10 ␮L of urine in a 96-deep-well plate. After
500 ␮L of distilled water was added to each well, the plate
was thermosealed and vortex-mixed. The plate was then
placed in a WatersTM 2795 Alliance HT LC system, and 5
␮L of each sample was injected on a SecurityGuard SCX
cation-exchange column [4.0 ⫻ 3.0 mm (i.d.); Phenomenex] attached to a Waters Atlantis® C18 column [50 ⫻ 2.1
mm (i.d.); 5 ␮m bead size]. Cystine was eluted from the
column by a stepwise gradient of 75% mobile phase A, 5%
B, and 20% C at 0 min; 5% A, 5% B, and 90% C at 0.2 min;
and 75% A, 5% B, and 20% C at 0.4 min. Mobile phase A