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Journal of Clinical Microbiology, December 2001, p. 4362-4369, Vol. 39, No. 12
Service de
Microbiologie,1 Service de
Génétique Moléculaire,2
Service d'Immunologie,3 and
Service d'Hématologie
Oncologique,4 Institut Gustave-Roussy, 94805 Villejuif Cedex, France
Received 24 May 2001/Returned for modification 22 August
2001/Accepted 18 September 2001
A real-time PCR assay was developed to quantify human
cytomegalovirus (CMV) DNA in peripheral blood leukocytes (PBLs) of bone marrow transplantation patients. Unlike other teams, we quantified CMV
and the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene using a plasmid containing both sequences as an external standard. Tenfold serial dilutions of this plasmid yielded overlapping standard curves that allowed the quantification of CMV and GAPDH gene copies in
an efficient and accurate manner. Sequential blood samples (164 specimens) were collected from 16 patients. PBLs were tested by the
pp65 antigenemia assay and quantitative CMV and GAPDH gene PCRs. CMV
DNA was detected by PCR in 13 patients a mean of 15 days prior to the
appearance of antigenemia. The administration of anti-CMV drugs led to
a rapid decrease in the numbers of viral copies and positive nuclei.
Real-time PCR assay results correlated with those of the CMV pp65
antigenemia assay (P < 0.00001). The TaqMan assay
may be a useful tool for rapid quantification of CMV infection and for
monitoring of CMV reactivation in bone marrow transplantation recipients.
Human cytomegalovirus (CMV) is a
well-known cause of mortality in blood and bone marrow transplantation
(BMT) patients. Monitoring of CMV reactivation from latency is critical
for these patients. Prophylaxis against CMV disease with ganciclovir
(8, 9) or foscarnet (16) in asymptomatic BMT
recipients has been shown to dramatically reduce the incidence of CMV
disease (8, 17). As ganciclovir and foscarnet are
myelotoxic and nephrotoxic, respectively, full treatment is often
started at the time of documented CMV reactivation. The key to
efficient and effective management of CMV infection in these patients
is a test capable of rapidly monitoring and quantifying the presence of
CMV in the blood. This is particularly essential for the identification
of subjects at high risk of developing CMV disease, e.g., patients
receiving steroid or immunosuppressive compounds for accelerated
graft-versus-host disease and also for the application and monitoring
of preemptive antiviral therapeutic strategies. The CMV assays
presently available and frequently used in this setting include shell
vial culture (7), the CMV antigenemia assay
(2), PCR for CMV DNA (3), hybrid capture assay for quantitation of CMV DNA (10), and detection of
CMV RNA by nucleic acid sequence-based amplification (1).
Among white blood cells, peripheral blood leukocytes (PBLs) are the
main CMV carriers during active CMV infection. The detection of CMV
antigenemia in PBLs has been shown to be an early marker of CMV
infection. A monoclonal antibody is used to detect pp65, the CMV lower
matrix phosphoprotein in PBLs, and this test is widely used to monitor
BMT recipients. Although a correlation was found between the number of
pp65-positive PBLs and the development of clinical symptoms, this
method (6, 20) poses a number of problems. It is difficult
to perform before engraftment because the number of leukocytes is
limited and false-negative results are obtained due to the poor
sensitivity of the technique and the weak expression of the pp65
antigen in white blood cells in some patients who develop CMV
disease (18). Alternatively, quantitative PCRs based on
TaqMan technologies for detection of CMV reactivation after BMT have
been investigated (14, 22), but to date, none of the
techniques described have been adequately standardized since the
numbers of CMV copies were never normalized by quantification of a
housekeeping gene.
The aim of our study was to design a quantitative PCR-based assay
capable of quantifying the CMV load in PBLs by two independent, quantitative PCR methods. The first PCR technique measured the CMV
genome copy number by using a target sequence located in the UL83 gene,
which codes for pp65. The second PCR technique quantified the
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene in order to
normalize the CMV DNA loads in the samples. The originality of our
study is that we used a plasmid (Pic19Rpp65/GAPDH) that contains both
CMV and GAPDH DNA fragments to quantify CMV DNA. We used these tools to
assess the usefulness of real-time automated PCR as a quantitative,
highly reproducible, and sensitive method for the detection of CMV DNA
in PBLs and to evaluate the extent to which it was correlated
with the CMV antigenemia assay in BMT recipients.
Patients and samples.
Sixteen patients who underwent BMT
between April 1998 and March 2000 were examined. Antigenemia assays
were performed in real time, and the results were used to determine
each patient's treatment. CMV DNA was retrospectively quantified in
PBLs in patients with either symptomatic or asymptomatic infection. In
the present study, 164 blood samples for the CMV antigenemia assay and
CMV DNA detection were drawn twice a week from the onset of aplasia
until the third month posttransplantation. Thirty-three additional
samples from BMT recipients were used to compare both methods. Thus,
the comparison of real-time PCR and the pp65 antigenemia assay was
performed with 128 PBL samples which were positive by PCR and/or by the antigenemia assay. According to our present CMV prevention strategy, between 25 and 35 days post-BMT, patients underwent a systematic bronchoscopy to obtain bronchoalveolar lavage (BAL) fluid samples for
CMV cultures (4, 11). All patients received prophylactic acyclovir at a dose of 10 mg/kg of body weight every 8 h from day
Sample preparation.
To prepare leukocytes, cells were
separated by sedimentation of 3 to 5 ml of a heparinized blood sample
in a 6% dextran solution over 20 min, centrifuged at 220 × g for 10 min at 20°C, and washed once with
phosphate-buffered saline (PBS). The resulting pellet was washed with 2 ml of erythrocyte lysing solution for 2 min and washed once with PBS.
Aliquots of 200,000 leukocytes were used for the pp65 antigen test, and
106 cells were used for DNA extraction, performed
with a QIAamp Blood mini-kit (Qiagen, Valencia, Calif.). The DNA
absorbed on the spin column was eluted with 50 µl of DNase-free
distilled water and was then submitted to PCR. The concentration of the
extracted DNA was quantified by spectrophotometric measurement at a
wavelength of 260 nm.
CMV antigenemia assays.
The CMV antigenemia assays
(15) were performed by indirect immunofluorescence
detection of pp65 (65 to 68 kDa), the human CMV internal matrix
phosphoprotein in PBLs, by standard procedures (CINA kit; Argene
Biosoft, Varilhes, France). Briefly, for each patient three cytospin
slides with 200,000 cells per glass slide were prepared. The
PBLs were fixed and permeabilized to allow subsequent detection of the
CMV pp65 antigen. The presence of the CMV pp65 antigen was detected
with the 1C3/AYM-1 antibody cocktail and was visualized with a specific
secondary antibody. The number of CMV antigen-positive cells was
counted on duplicate stained slides, and the results were
reported as the number of positively staining cells per 200,000 leukocytes.
Real-time quantification PCR.
The sequences of the PCR
primers and that of the probe used to quantify CMV were selected from
the phosphorylated matrix protein (pp65) gene (locus HSPPBC; GenBank)
with Primer Express software (Perkin-Elmer Biosystems, Foster City,
Calif.). The sequences of the forward and reverse primers were
5'-GCAGCCACGGGATCGTACT and
5'-GGCTTTTACCTCACACGAGCATT, respectively. The TaqMan probe selected between the primers was fluorescence labeled at the 5' end
with 6-carboxyfluorescein (FAM) as the reporter dye and at the
3' end with 6-carboxytetramethylrhodamine (TAMRA) as the quencher (5'-FAM-CGCGAGACCGTGGAACTGCG-TAMRA). The PCR product was detected as an
increase in fluorescence with the ABI PRISM 7700 instrument (Perkin-Elmer Biosystems). PCR was performed with 25 µl of TaqMan Universal PCR master mixture (Perkin-Elmer Biosystems), each of the
primers at a concentration of 400 nM, 100 nM TaqMan probe, and 100 ng
of DNA in a total volume of 50 µl. PCR was performed in 96-well
microtiter plates under the following conditions: after 2 min at 50°C
and 10 min at 95°C, the samples were submitted to 45 cycles, with
each cycle consisting of a step at 95°C for 15 s,
followed by a step at 60°C for 1 min, for both CMV and GAPDH amplification. The human genomic sequence was quantified with PCR
primers and the TaqMan probe that recognized the GAPDH gene. The
upstream and downstream primer sequences were
5'-CTCCCCACACACATGCACTTA and
5'-CCTAGTCCCAGGGCTTTGATT, respectively, and the fluorogenic probe located between the PCR primers and labeled with VIC and TAMRA
was synthesized by PE Biosystems
(5'-VIC-AAAAGAGCTAGGAAGGACAGGCAACTTGGC-TAMRA). The GAPDH
gene PCR was performed under the same PCR conditions described above
for the CMV PCR. A plasmid containing both targeted sequences was used
as a standard in the present study. This plasmid (Pic19Rpp65/GAPDH)
contained a 448-bp DNA fragment derived from the pp65 gene and a 538-bp
fragment derived from the human GAPDH gene, which were generated by
conventional PCR with Pfu polymerase (Stratagene, Amsterdam,
The Netherlands). A standard graph of the cycle threshold
(CT) values obtained from serial dilutions (10 to 104 copies/well) of the plasmid was
constructed for both CMV and the GAPDH gene. The 10-fold dilutions of
the plasmid were concocted with a solution of salmon sperm DNA as a
carrier at a final concentration of 100 ng of DNA per sample. The
CT values from unknown samples were
plotted on the respective standard curves, and the number of CMV genome
copies per 200,000 leukocytes was calculated with Sequence Detection
System software (version 1.6.3; Perkin-Elmer Biosystems). Each sample
of DNA extract and serial dilutions of the plasmid were analyzed in
duplicate. To control for cross-contamination, a sample consisting of
distilled water was also submitted to the DNA extraction procedure, and
the resulting extract was amplified in duplicate. Samples were
considered negative if the CT values exceeded 45 cycles.
TaqMan PCR design: PCR specificity, sensitivity, and
reproducibility.
Our goal was to design a quantitative PCR-based
assay capable of quantifying the CMV load in human cells. Two
independent, quantitative PCR methods were used and performed with two
separate aliquots of the same DNA extract. The CMV TaqMan PCR is based on the amplification of a 159-bp region of a sequence located in the
UL83 gene, which codes for the lower matrix protein detected in the
pp65 antigenemia test. A nonvariable region of the pp65 gene, common to
strains AD149 and Towne, was chosen. The GAPDH gene PCR is based on the
amplification of a 99-bp region of human genomic DNA. In order to
confirm the specificity of this assay, several virus strains were
tested. No cross-reactivity between CMV and herpes simplex virus types
1 and 2, varicella-zoster virus, Epstein-Barr virus, and human
herpesviruses 6 and 8 was observed (data not shown).
0095-1137/01/$04.00+0 DOI: 10.1128/JCM.39.12.4362-4369.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Quantification of Human Cytomegalovirus DNA in Bone Marrow
Transplant Recipients by Real-Time PCR
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
1 until discharge from the BMT unit or the beginning of
anti-CMV therapy, initiated at the time of documented CMV reactivation on the basis of the results of the pp65 antigenemia assay. For all
patients, prophylaxis against graft-versus-host disease consisted of
cyclosporine A on the day before the graft and a short course of
methotrexate on days 1, 3, and 6 posttransplantation.
RESULTS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
10 copies per well; and the detection rates were 60 and 40% for the GAPDH gene and CMV PCRs, respectively, when the copy
number was 1 copy per well, which is in agreement with the values that
can be estimated from the Poisson probabilities. As shown in Fig.
1A, B, and D, the amplification yield and
detection rates were comparable when plasmid dilutions were submitted
to both PCR techniques, with CT values of
38.9 ± 0.67 (values are means ± standard deviations) and
38.71 ± 0.69 for the amplification of 10 copies of the GAPDH gene
and CMV, respectively. In this test of intra-assay variation, the
coefficients of variation (CVs) were 0.17, 0.13, 0.32, and 1.71% for
104, 103,
102, and 10 copies/well, respectively, for the
GAPDH gene PCR and 0.25, 0.55, 1.03, and 1.79% for
104, 103,
102, and 10 copies/well, respectively, for the
CMV PCR. As shown in Fig. 1C, the linear correlations between the
CT value and the logarithm of the DNA copy
number were identical for both PCRs. The slopes were 3.70 and 3.62
for the GAPDH gene and CMV PCRs, respectively, and the correlation
coefficient was identical for both PCRs
(R2 = 0.9968).
View larger version (45K):
[in a new window]
FIG. 1.
Amplification plots obtained with the control plasmid
for GAPDH gene (A) and CMV (B) PCRs ( 
Rn, normalized reporter
fluorescence signal). Serial 10-fold dilutions with 104 to
10 copies per reaction well were amplified for 45 cycles, and
amplifications were repeated four times for each dilution. (C)
Standard curves for CMV and GAPDH gene real-time PCRs.
CT values were plotted against the
normalized fluorescence signal. The correlation coefficient was 0.9968 for both PCRs, and the slopes were 3.7 and 3.6 for the GAPDH gene
and CMV PCRs, respectively. (D) After amplification of 104
to 0 copies of the plasmid, the PCR products were loaded onto an
ethidium bromide-stained 2% agarose gel.
Patient characteristics.
Patient characteristics are
summarized in Table 1. There were 10 females and 6 males, with a median age of 34 years (age range, 8 to 52 years). Conditioning for allogeneic BMT consisted of total body
irradiation (n = 12) or busulfan (n = 3) combined with endoxan, melphalan, etoposide, and/or cytarabine. One
patient received a reduced conditioning regimen with busulfan, endoxan, anti-T globulin, and fludarabine. Fifteen patients were seropositive for CMV before BMT, and they received a graft either from seronegative donors (n = 4) or from seropositive donors
(n = 11). Patient 16 and his donor were seronegative.
The BAL fluid samples of two patients were positive for CMV, and the
patients were immediately prescribed a 14-day course of intravenous
ganciclovir at a dose of 10/mg/kg/day.
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PCR quantification of CMV DNA and comparison with pp65 antigenemia
assay.
We tested 197 PBL samples from BMT recipients to evaluate
the CMV DNA TaqMan assay with clinical specimens. CMV DNA was
quantified in parallel with the GAPDH gene in PBL samples in order to
determine the amount of cellular DNA input into each reaction mixture.
In our study, the concentration of the extracted DNA was quantified by
spectrophotometric measurement at a wavelength of 260 nm; this measurement was only an approximation of the DNA concentration in each
sample, i.e., 330 copies per ng of genomic DNA, in theory. The mean
value of the GAPDH gene copy number for the 197 PBL samples tested was
87,974 copies (range, 24,779 to 296,295 copies), which corresponded to
2.6 times (range, 0.75 to 8.9 times) the GAPDH gene copy number
expected for the amount of cellular DNA input into the reaction
mixture. Of the 197 samples tested, 69 were negative by both
techniques. The 128 samples that were PCR positive were classified into
three groups according to the results of the pp65 antigenemia assay.
Samples in group 1 (n = 66) were negative for pp65
antigenemia, samples in group 2 (n = 37) had <10
pp65-positive cells, and samples in group 3 (n = 25)
had 10 pp65-positive cells. As shown in Fig.
2, the samples in group 3 had
significantly higher CMV DNA copy numbers than the numbers in the
pp65-positive samples in group 2 (P = 0.001 by the
Student test) and the pp65-negative samples in group 1 (P < 0.0001 by the Student test). The mean CMV DNA
copy number per 2 × 105 PBLs was 138 (range, 4 to 3,444) for samples in group 1, 296 (range, 7 to 1,770) for
samples in group 2, and 2,164 (range, 36 to 14,446) for samples in
group 3. Furthermore, a statistically significant correlation was
observed between the CMV DNA copy number and the number of
pp65-positive cells (n = 128; r = 0.425; P < 0.00001 by the Spearman rank test). Figure
3 shows on a logarithmic graph the
results of a comparison of both techniques in which samples which were
negative for CMV DNA or pp65 antigenemia (n = 66 samples from group 1) were not included.
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Analysis of BMT patients.
The quantification of CMV DNA was
performed retrospectively with 164 samples from 16 BMT recipients
tested for the pp65 antigen, with a median number of 10 samples (range,
7 to 14 samples) available from each patient. Table
2 shows
the results of monitoring of CMV reactivation by the pp65 antigen test
and quantification of CMV DNA by PCR. As shown in Table 2, 42 of 164 samples (25.3%) were positive for CMV DNA and pp65 antigen detection,
while 69 of 164 samples (41.5%) were negative by both tests.
Fifty-three pp65 antigen-negative samples (32%) were PCR positive.
After BMT, the CMV DNA PCR was positive for 13 of 15 patients a mean of
15 days before the pp65 antigenemia assay was positive (84 ± 69 and 69 ± 65 days for the pp65 antigenemia assay and PCR,
respectively). For two patients (patients 3 and 11) the two tests were
positive simultaneously. Patient 16, who had a negative CMV serology,
as did his donor, was negative by both techniques. All 15 patients received antiviral therapy, which was started on the day that positivity for pp65 antigenemia was documented. A single course of
ganciclovir therapy led to marked decreases in CMV DNA loads and levels
of antigenemia, but significant differences were observed among
patients. Eight patients (seven CMV-positive patients with CMV-positive
donors and one CMV-negative patient with a CMV-positive donor)
responded promptly to a single 14- to 59-day course of therapy, and
both the pp65 antigenemia assay and the CMV DNA PCR became negative
after ganciclovir therapy. For six patients (patients 3, 6, 7, 8, 9, and 11), both techniques became negative on the same day posttreatment.
For patients 10 and 12, the pp65 antigenemia assay was negative before
negativity by the CMV DNA PCR, with CMV DNA still being detectable 14 and 21 days posttreatment, respectively. Seven patients exhibited
asymptomatic CMV reactivation after the first course of ganciclovir
therapy, and a second pp65 antigenemia and/or CMV DNA peak was
observed. Among these seven patients, the second peak was observed by
both techniques in five patients (patients 1, 2, 4, 5, and 14) but only
by PCR in the other two patients (patients 13 and 15). In five patients
exhibiting CMV reactivation (patients 1, 2, 5, 13, and 15), CMV DNA was
still detectable 111, 151, 88, 73, and 69 days posttransplantation, respectively.
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DISCUSSION |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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The development of real-time PCR technology is a promising advance in the quantification of CMV DNA in clinical samples and will be useful for the monitoring of CMV reactivation in patients at high risk of developing CMV disease. In our study, we assessed PBLs for increased levels of CMV DNA replication, and an increased level of CMV DNA replication in PBLs was shown to be the first manifestation of CMV reactivation, which is consistent with previous findings (12). As PBLs were tested, the variations in the efficiency of the DNA extraction step and in the measurement of DNA levels by spectrophotometry could have major effects on the reproducibility of the results. Consequently, and as proposed by others, quantification of a housekeeping gene to normalize CMV loads is essential to guarantee highly reproducible PCR results (5, 14). Unlike other studies, we used a plasmid that harbors both sequences as an external standard to quantify CMV DNA and genomic DNA. We have shown that serial dilutions of this plasmid allowed the quantification of CMV and GAPDH gene copy numbers in an efficient and accurate manner, since the standard curves for CMV and GAPDH gene copy numbers were identical (Fig. 1C). Furthermore, our assay was highly reproducible, as indicated by the intra-assay and interassay CV values obtained with the standard plasmid. In our study, the intra-assay variation values were 0.17, 0.13, 0.32, and 1.71% for samples containing 104, 103, 102, and 10 copies, respectively, for the GAPDH gene PCR and 0.25, 0.55, 1.03, and 1.79% for samples containing 104, 103, 102, and 10 copies, respectively, for the CMV PCR. The interassay variations were less than 2% for DNA inputs of 104, 103, and 102 copies/well and less than 3.3% for DNA inputs of 10 copies/well for both the CMV and the GAPDH gene TaqMan PCRs. We consider that these variations will be negligible in clinical use.
The real-time PCR technique used in the present study allowed the quantification of CMV DNA over a wide dynamic range for both CMV and GAPDH gene amplification (10 to 104 and 10 to 105 copies of plasmid, respectively). Furthermore, we found a good positive correlation between the number of copies of CMV DNA and the GAPDH gene present and the number of cycles at which the amplification curves became steep using serial dilutions of plasmid Pic19Rpp65/GAPDH. In this 10-fold dilution range, we were able to accurately determine the number of copies of viral and genomic DNAs present in all samples used in our study.
In agreement with the results of other studies that used other TaqMan-based assays (5, 13, 19), we found a significant correlation between the results of the antigenemia assay and DNA copy numbers in PBL samples, but we nevertheless observed inconsistencies. Interestingly, 32% of the samples were pp65 antigenemia assay negative and PCR positive. Furthermore, samples with low levels of antigenemia and a large number of copies of CMV DNA were observed. Similar differences between the results of an antigenemia assay and quantitative PCR have been reported by others (5, 14, 21), but the clinical significance of large CMV DNA copy numbers with a low or a negative antigenemia assay result remains to be elucidated.
We have shown that PCR quantification of CMV DNA is more sensitive than the antigenemia assay for the monitoring of CMV reactivation in BMT patients. As shown in Table 2, CMV DNA was detected by PCR in 13 of 15 patients a mean of 15 days prior to the detection of antigenemia. All 15 patients were administered anti-CMV drugs when the antigenemia assay was positive. The antigenemia assay and the real-time automated PCR both showed rapid decreases in the numbers of viral copies and pp65-positive nuclei once treatment with ganciclovir was initiated. In our opinion, real-time PCR showed greater sensitivity than the antigenemia assay in reflecting the effect of ganciclovir since the samples became negative by the antigenemia assay before the quantification of CMV DNA in more than 50% of the patients. Furthermore, real-time PCR will permit more reliable screening than the antigenemia assay for patients who develop ganciclovir-resistant CMV, in whom CMV DNA replication is not inhibited.
All but two patients had detectable CMV DNA and pp65-positive nuclei in the absence of clinical manifestations of the disease. A factor that compounds difficulties in correlating the amounts of CMV DNA with the development of active disease is the widespread use of preemptive therapy. Two patients in whom CMV DNA was detected before antigenemia (by 7 and 37 days, respectively) developed active CMV-induced lung disease 29 and 34 days posttransplantation, respectively. Furthermore, in one patient (patient 6), the presence of infectious CMV in BAL fluid was detected at day 29 posttransplantation, whereas CMV DNA was quantified 7 days before the BAL. These findings support the use of quantitative PCR to assess when preemptive therapy should be initiated. Furthermore, the early detection of CMV DNA in PBLs should spare these patients from undergoing BAL, which is generally carried out between 25 and 35 days posttransplantation.
On the basis of our findings, we believe that real-time PCR promises to be an interesting alternative test to the antigenemia assay presently in use for the monitoring of CMV disease in BMT patients. We are therefore conducting a prospective study with a large patient population in which half of the dose of preemptive antiviral therapy is administered when CMV reactivation is documented by the CMV TaqMan PCR and until CMV DNA is no longer detectable. This approach will avoid the myelotoxicity and the nephrotoxicity caused by the drugs usually prescribed (ganciclovir and foscarnet, respectively) and should therefore significantly improve the clinical conditions of these patients.
In summary, we have presented an accurate and rapid PCR assay for the quantification of CMV DNA which may prove useful for routine clinical testing. We have shown that the standardization of the technique requires the amplification of a cellular gene to monitor the efficiency of the reaction. Since both CMV- and GAPDH gene-specific probes are labeled with two different fluorogenic dyes (FAM and VIC, respectively), a multiplex PCR technique will be developed in order to decrease the cost of the TaqMan PCR.
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ACKNOWLEDGMENTS |
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We are grateful to all the staff of the Microbiology Laboratory of the Institut Gustave-Roussy and to L. Saint Ange and M. Mackenthun for editing. We thank E. Dussaix and A.-M. Roque for providing us with herpes simplex virus type 1 and 2, varicella-zoster virus, Epstein-Barr virus, and human herpesvirus 6 and 8 DNAs.
This study was supported by a CRC (Contrat de Recherche Clinique) grant from Institut Gustave-Roussy.
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FOOTNOTES |
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* Corresponding author. Mailing address: Laboratoire de Microbiologie Médicale, Institut Gustave-Roussy, 39 rue Camille Desmoulins, 94805 Villejuif Cedex, France. Phone: (33-1) 42 11 51 93. Fax: (33-1) 42 11 53 13. E-mail: grisceli{at}igr.fr.
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Discussion
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Journal of Clinical Microbiology, December 2001, p. 4362-4369, Vol. 39, No. 12
0095-1137/01/$04.00+0 DOI: 10.1128/JCM.39.12.4362-4369.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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