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TRANSFUSION MEDICINE
From Gambro BCT, Lakewood, CO; University of Colorado
Health Sciences Center, Denver; Eastern Virginia Medical School,
Norfolk; American Red Cross, Mid-Atlantic Region, Norfolk, VA; and
Bonfils Memorial Blood Center, Denver, CO.
This study examined the effectiveness of 3 leukocyte-reduction (LR)
methods in depleting the residual level of cytomegalovirus (CMV) in
blood products measured by quantitative polymerase chain reaction
(QA-PCR). At 2 locations over 3 allergy seasons, apheresis platelets
and whole blood were collected from 52 healthy CMV seropositive subjects having an elevated titer of CMV DNA (median = 2400 genome equivalents [GE]/mL) resulting in 32 evaluable LR apheresis
platelets, 31 filtered platelets from whole blood, and 31 filtered red
blood cells (RBCs) from whole blood. Leukoreduction by apheresis and filtration resulted in substantial reduction of detectable CMV DNA
levels with 99.9% of the LR products expected to have less than 500 GE/mL of CMV DNA. No difference was found between methods (P = .52). CMV genomic leukocyte subset localization was
determined by QA-PCR of fluorescence-activated cell sorter
(FACS)-sorted peripheral blood from 20 seropositive subjects
(n = 10 > 100 GE/mL, n = 10 QA-PCR negative). CMV was detected
in monocyte (13 of 20) and granulocyte (3 of 20) fractions. Presence of
competent virus in QA-PCR positive (> 100 GE/mL) peripheral blood
samples was verified with 4 of 19 subjects positive in shell vial
assay, and 8 of 18 positive for CMV gene products (messenger RNA). We
observed a seasonal DNAemia variation in seropositive subjects. CMV
seropositive subjects (n = 45) entered into longitudinal monitoring
in March/April 1999 were QA-PCR negative at baseline. Subjects
converted to a positive QA-PCR coincident with increased seasonal
allergen levels (Norfolk 15 of 18 evaluable in 43.4 ± 9.48 days;
Denver, 16 of 23 evaluable in 96 ± 26.3 days). These data
demonstrate effective reduction of CMV load by LR during periods of
DNAemia in CMV seropositive subjects.
(Blood. 2001;97:3640-3647) Transfusion-transmitted cytomegalovirus (CMV)
infections cause only mild illness in individuals with normal immune
systems, but can cause serious and even fatal illness (including
retinitis, gastroenteritis, and interstitial pneumonitis) in the
seronegative, immunocompromised recipient.1,2 To avoid
these serious complications, it is recommended that seronegative
immunocompromised patients receive blood components from seronegative
donors.3 Typically, the majority of the blood donors in a
geographic region are seropositive for CMV, which makes providing CMV
seronegative blood components at times a difficult logistical task.
Because it is believed that latent CMV resides in the white blood cells
(WBCs), an alternative to using seronegative donors has been to remove
most of the leukocytes from transfused products.4 A number
of studies have suggested that reduction in the total number of
leukocytes in blood products (leukocyte-reduction [LR]) below
1 × 107 to 1 × 106 would reduce the risk
of transmission of CMV to a level comparable to that of seronegative
blood products.5,6 Such levels of LR have been achieved
either by filtration7 or during collection of apheresis
platelets using a cell separator.8
Some investigators have suggested that various methods of LR result in
different distributions of WBC subsets in the final product. Based on
the assumption that latent CMV is only in a particular cell type, they
further extrapolate these data to suggest that different methods of
preparation will result in varying effectiveness for reduction of the
viral load presented by the blood donor.9 Residual WBCs in
leukocyte-reduced products are rare events with typically fewer than
100 cells/mL. The seemingly simple process of obtaining an accurate WBC
count in these products has historically been difficult. Further
isolation and subtyping of these cells presents many technical
challenges, such as isolating WBCs without cell loss and observing
enough events to make accurate estimates. Inferences regarding CMV
safety from these data require additional biologic assumptions
regarding the exclusive location of latent or active virus. We wished
to determine if there is a difference in the effectiveness of LR
methods in rendering blood components CMV safe, and believed
quantitative polymerase chain reaction (QA-PCR) offered the most
sensitive measure of low levels of virus without the pitfalls of the
previously mentioned assumptions.
Study subjects
Blood products
Products collected in Denver were shipped overnight with 22°C gel packs from Lakewood to Norfolk for CMV QA-PCR. Subject peripheral blood samples used for CMV QA-PCR screening and reverse transcription-PCR (RT-PCR) were collected in EDTA tubes and shipped overnight at 4°C. Blood products and samples were held at 4°C 0 to 3 days before DNA extraction. WBC subset isolation and testing for CMV genome To determine which WBC subsets carried the CMV genome, PCR+ and PCR subject samples from Norfolk
during the spring of 1998 were separated by density gradient separation
using OptiPrep density gradient media (Nycomed Pharma AS, Oslo, Norway)
into monocyte, lymphocyte, and granulocyte fractions. The separation
was done according to the protocol recommended by the manufacturer. The
enriched monocytes (present in the top fraction of the gradient) were
further purified by FACS (BD, San Jose, CA) based on CD14 markers to
remove the small amount of contaminating lymphocytes. The lymphocyte
fractions (present in the middle part of the gradient) were further
purified by FACS using CD3, and CD4 markers to isolate T cells, and
CD19 marker to isolate B cells (BD; Diatec AS, Oslo, Norway).
Granulocytes present in the pellet fraction were used directly for
further analysis. Analysis of the latter fraction indicated that
contamination with CD14+ cells was less than 0.5% and
contamination with lymphocytes (CD3, CD4, CD19 combined) was less than
3%. One million (1 × 106) cells were used from the
isolated granulocytes, monocytes, B lymphocytes, and T lymphocytes for
detection of the presence of CMV genome by QA-PCR and by RT-PCR for
expression of viral gene products p72 and viral DNA polymerase.
QA-PCR DNA preparation from blood or blood products. DNA was extracted from 100µL peripheral blood or 1 × 106 cells from the various cell fractions by lysing cells, solubilizing proteins with 1 mL DNAzol (Gibco-BRL, Rockville, MD) and precipitating with 0.5 mL ethanol according to the manufacturer's recommendation. DNA was washed twice with 70% ethanol, dried, and resuspended in water for use in PCR reactions. Leukocyte-reduced platelets and RBCs were prepared for QA-PCR by fully extracting DNA from 60 mL using a silica-based method. Briefly, to 60 mL product was added 32.0 g guanidine-HCl, 1.2 mL 2M Tris-HCl, pH 7.4, 0.6 mL 0.5M EDTA, and 120 µL 2-mercaptoethanol. After dissolution of the guanidine-HCl, the mixture was incubated for 10 minutes at 37°C to solubilize membrane proteins. Prewashed silica (100 µL of a 30% mix in water) was added and mixed for 15 minutes. The silica was separated from the solution by centrifugation, washed 3 times in 4 M guanidine-HCl, washed twice in ice-cold Tris-HCl, pH 7.4/50% ethanol (pH 7.4, 5 mM EDTA), and dried. Water (100 µL) was added to the silica and the bound DNA was eluted by incubation at 57°C for 5 minutes. The silica was removed by centrifugation and the DNA was precipitated with sodium acetate and isopropanol, washed with 70% ethanol and dried, and finally resuspended in 100 µL water for PCR.CMV QA-PCR assay.
Quantitative PCR was performed by amplifying a 240-base pair (bp)
region of CMV DNA-polymerase gene using primers 5'-GCT ATG TTT CAG ATG
TCG CCG CC (5'-biotin), 5'-CCC ACC TCG GGC TCA AAC AC (Midland
Certified Reagent, Midland, TX). One microgram of DNA per reaction tube
was amplified using the AmpliTaq Gold enzyme (PerkinElmer, Norwalk,
CT). The conditions were: initial melt 94° for 6 minutes, followed by
39 cycles melting at 94°C for 20 seconds and annealing at 58°C for
30 seconds and extension at 72°C for 40 seconds. Amplification
products were resolved by acrylamide gel electrophoresis, developed
with ethidium bromide, and visualized with an imaging system (Stratagen
Eagle Eye II Camera System, La Jolla, CA). The image was stored
as a TIFF file and the intensity of the reaction products were analyzed
using SigmaGel (SPSS Science, Chicago, IL). CMV+ or
CMV
Because of the very low levels of residual WBCs in the blood products (typically < 3 WBC/µL), a positive control for DNA amplification was run with each sample. A 430-bp region of the hemachromatosis gene was amplified as above using primers 5'-TCC TGG CAA GGG TAA ACA GAT CC and 5'-CTC CTC AGG CAC TCC TCT CAA CC (5'-biotin). Resolved amplification products were identified using the probe 5'-CAA GGA GTT CGA ACC TAA AGA CGT ATT GCC CA. The assay was completed as described for CMV genome detection. The sensitivity of detection of the hemachromatosis gene was 50 copies per PCR reaction. Determination of the detection limit of
QA-PCR.
Because no cell lines containing latent CMV genomes are in existence,
and independent quantitation of CMV genomes in infected cells is not
possible, the detection limits for viral genomes in these preparation
and assay methods were determined using blood products spiked with
Namalva cell line containing 2 copies of Epstein-Barr virus (EBV) per
cell. The primers used for this assay were 5' TAT GAC AAA GCC CGC TCC
TAC CT and 5' GGG AAT ACA CGG CTT TTA ATA CG and the size of the PCR
product was 236 bp. The sensitivity of EBV PCR was similar to the CMV
PCR as was determined in previous studies. This method allowed us not
only to determine the lowest viral copy numbers to be detected but also
indicated that we can isolate DNA from fewer than 500 cells diluted in
60 mL blood product. Spiked RBCs and platelets were extracted using the
silica protocol followed by QA-PCR using primers for EBV. The lower
detection limits of 100 GE/mL (genomic equivalent) for peripheral blood
and 4 GE/mL for the blood products were observed (Figure
2). However, because detection of CMV
genome and EBV genome is done with different primers, the possibility
exists that the sensitivity corresponding to the EBV genome cannot be
directly adapted to the detection of CMV genome. In a recent
multilaboratory blinded study our laboratory could detect and
accurately quantitate 10 genome copies of CMV in duplicate samples
indicating that the QA-PCR adapted in our laboratory can reproducibly
detect low copy numbers of CMV (manuscript submitted for
publication).
RT-PCR Cell pellets (1 × 106 to 5 × 106) from whole blood or cell fractions were resuspended in Trizol reagent (Gibco-BRL) and total RNA was purified according to the manufacturer's directions. Polyadenylated messenger RNA (mRNA) was isolated from the purified RNA by binding the mRNA to Instant mRNA Capture Disc (Trevigen, Gaithersburg, MD) according to the manufacturer's recommendation. Bound mRNAs were eluted with 50 µL Rnase-free water for use in the RT-PCR assay. The assay was performed with 10 µL of the mRNA preparation (corresponding to 1 × 106 cells) using the EZ-RT-PCR kit (PerkinElmer). The tubes were incubated at 62°C for 20 minutes to convert RNA to DNA; then amplification was performed by cycling 36 times to 94°C for 0.3 minutes, annealing at 58°C for 0.4 minutes, and elongating at 72°C for 0.5 minute. Amplification products were resolved by acrylamide gel electrophoresis, developed with ethidium bromide, and visualized with an imaging system (Stratagen Eagle Eye II Camera System).CMV culture To determine if the detection of CMV genome was indicative of the presence of infectious virus, samples from positive donors during the spring of 1999 were evaluated via shell vial assay. Although primary cell line cultures of various foreskin fibroblast are frequently used in some cell vial assays, we found that MRC-5 fibroblast cells can be equally well infected with CMV strains from various patients to be used in this assay. Therefore, the fibroblast cell line MRC-5 (American Tissue Culture Collection, Rockville, MD) was used in our shell vial assays. Cells were overlaid in triplicate with buffy coat purified from 1 mL of subject peripheral blood sample and incubated at 37°C, 5% CO2 for 10 days. Cultures were checked for infectivity at days 3 and 10 by staining the cells with a monoclonal antibody against the CMV p72 antigen and detection of p72 mRNA expression by RT-PCR.Cytokine assays Plasma was separated within 30 minutes from peripheral blood samples collected in EDTA and frozen at 70°C. Quantitative sandwich enzyme immunoassays were performed on thawed plasma for interleukin (IL)-2, IL-4, and IL-10 and tumor necrosis factor- (TNF-) (R&D Systems, Minneapolis, MN) according to the manufacturer's instructions.
Pollen data Local pollen data were kindly provided by Dr Alpha A. Diallo, Department Public Health, Norfolk, and obtained from the American Academy of Allergy, Asthma, and Immunology Web site for Denver (data collected at National Jewish Medical and Research Center).Statistical methods The primary end point, CMV genome residual in leukocyte-reduced blood products, was evaluated as a dichotomous outcome variable (detectable or nondetectable at 4 GE/mL) for those procedures where there was detectable CMV genome in the subject before collection. Matched subject effects for the 3 preparation methods were evaluated by the McNemar 2 test. Logistic regression of outcome
variable on preparation methods was performed with global testing of
equivalence of the preparation methods. As secondary analyses,
estimates of the probability of detection an global equivalency tests
were conducted using logistic regression at detection thresholds of 50 and 80 GE/mL in the products.
As additional secondary analyses, residual WBCs in the products were evaluated as a log-normal distribution as described by Dumont and colleagues.14 The effect of residual WBCs on final genome concentration in the end products was determined by logistic analysis that included preparation methods as predictor variables. The association of subjects' reported allergy condition with the level of peripheral blood CMV genome was evaluated with a general linear regression model of the logarithm of peripheral blood CMV regressed on questionnaire responses. The Fisher exact test was applied to the differences in questionnaire responses between sites. All tests were performed at an alpha of 0.05.
During February and March 1998, 148 CMV-seropositive blood donors
at the American Red Cross, Norfolk. Zero versus an anticipated 35% to
50% of these donors were positive for CMV DNA by QA-PCR. Because of
previous local observations of season association with CMV DNAemia, the
screening was suspended until April, whereupon the CMV positivity
increased to 39 of 40 subjects (95%, median = 4200 GE/mL,
range = < 100-1 000 000 GE/mL). Nine of these subjects with
median screening levels of 15 800 GE/mL were then entered into phase 2 (blood product collection). Logistical delays resulted in up to a
30-day interval from the time of screening until the first collection,
whereupon many of the subjects' levels had dropped to below detection
level. Noticing the apparent seasonal association of this reactivation,
we reinitiated screening during the fall of 1998 in Denver and Norfolk.
Although reactivation was observed during the fall mold seasons at both
locations, the absolute peak levels of CMV DNA were lower
than observed in the spring with a rapid falloff over 2 weeks.
Therefore, in the spring of 1999 enrollment was increased in both
locations, gathering longitudinal data on subjects over a long calendar
period. In Norfolk, 20 CMV-seropositive subjects were entered into
weekly monitoring (phase 1) between March 2 and 30, 1999. Two subjects
withdrew from the study and 15 of 18 converted to a positive CMV
DNAemia in a mean of 43.4 days (SD = 9.48, range = 22-57 days).
Twelve positive subjects were entered into the collection phase (phase
2). Subjects were followed for a maximum of 65 days, through May 5. In
Denver, 25 CMV-seropositive subjects were entered into phase 1 between
March 15 and April 23, 1999, with initial monitoring every other week changing to weekly as the Denver tree pollen season approached. Of
these, 2 subjects withdrew and 16 of 23 converted to a positive CMV
DNAemia in a mean of 96 days (SD = 26.3, range = 0-120 days). Thirteen positive subjects were entered into phase 2. Subjects were
followed for a maximum of 120 days, through July 7. Figure 3 shows the maximum CMV DNA titers
observed for subjects throughout the monitoring and collection phases.
Norfolk subjects reactivated during April and Denver subjects during
June/July, both following the local tree pollen rise.
To determine the WBC subtype localization of the CMV, we identified 10 CMV DNA positive and 10 negative subjects from the April-May 1998 Norfolk screening. During a subsequent visit, a second sample was taken
and WBC subsets isolated as described in "Patients, materials, and
methods." The CMV was detected in monocytes (CD14) in 13 of 20 samples; even for those samples that had an original whole blood level
of less than 100 GE/mL. CMV was detected in the density gradient
granulocyte fraction for 3 subjects (Table
1).
Aggregate CMV QA-PCR levels are shown for phase 2 treatments in Figure
4. Reactivated subjects in the spring of
1999 were entered into phase 2 within 7 days of a positive screening
result. Subject preprocedure CMV DNA levels in phase 2 were 50 to 100 times above the limit of detection of the assay. Blood products collected were within the typical ranges of volume and cellular content
(Table 2). Residual WBC in the products
were: filtered RBC median = 7.1 × 104, 98.8% less
than 1 × 106 (n = 51); filtered platelets
median = 7.9 × 103, 99.9% less than
1 × 106 (n = 51); apheresis platelets
median = 4.0 × 104, 85% less than
1 × 106 (n = 52). Seven of the apheresis platelet
procedures were flagged as nonleukocyte reduced by the equipment. Note
that flagged procedures in clinical practice would not be regarded as
CMV safe. Procedures indicated by the apheresis equipment as leukocyte
reduced resulted in apheresis platelet WBC residuals
median = 3.1 × 104, 99.8% less than
1 × 106 (n = 45). However, all apheresis procedures
(n = 52) were included for the CMV analysis.
Phase 2 collections with the subject predonation CMV DNA level more
than 100 GE/mL were selected for preparation method effect analyses
resulting in 32 of 52 evaluable apheresis products, 32 of 51 evaluable
filtered platelets, and 32 of 51 evaluable filtered RBC. The effect of
matched preparation methods with one subject undergoing all treatment
arms was evaluated by the McNemar
Pooled Norfolk and Denver subject CBC at baseline and immediately
before phase 2 collections are shown in Table
4. There were no significant changes in
baseline numbers taken in February and those at the height of the CMV
DNAemia. A small increase in mean lymphocyte count was observed
(P = .037). Other changes, which may have been indicative
of an allergic response, such as eosinophilia, were not seen.
Subjects entering the collection phase in the spring of 1999 were presented with a questionnaire, which asked about various aspects of allergic status and treatment. There were no differences between the Norfolk and Denver groups for known allergies or receipt of allergy shots (P < .16). No Denver subject and 33% of Norfolk subjects were being seen by an allergist (P = .039). None of these categories were significantly correlated with the maximum level of CMV titer observed in the subject (P > .5). To determine if reactivation may be caused or associated with increased
systemic cytokine levels, a panel of cytokines (IL-2, IL-4, IL-10, and
TNF- Determination of competent virus via mRNA detection and tissue culture
are shown in Table 5. Messenger RNA was
detected in 8 of 19 subjects. Four of 19 subjects had positive tissue
culture by day 10. Both transcription and positive culture observations were associated with higher levels of genome copies.
In this study, we have observed a seasonal variation in the level
of CMV DNAemia that is associated with increases in environmental allergens, most notably pine tree pollen. This effect has been previously observed in the Norfolk area, but in this study confirmed in
2 locations over 2 allergy seasons, with CMV DNAemia varying from 0%
to 100% in normal healthy CMV seropositive subjects. Various investigators have reported a low frequency of CMV DNAemia in serologically positive blood donors (Bitsch and
coworkers15 0 of 100 [0%], Urushibara and
colleagues16 0 of 155 [0%], Smith and
coworkers17 7 of 86 [8%], and Krajden and
associates18 0-8 of 101 [0%-8%]). More recently,
Larsson and coworkers19 observed 41.4% (60 of 145) CMV
DNAemia in seropositive blood donors. In a small cohort followed over
time, Larsson also found that seropositive donors initially negative
for CMV DNA converted to positive. The variety of specific preparation
and amplification methods used in these studies may explain some of the
variations among the groups. Our data show latent CMV may be at very
low levels (< 100 GE/mL) in the peripheral blood of seropositive, healthy individuals, then reactivated resulting in extremely high, transient titers of CMV DNAemia (> 1 000 000 GE/mL). The triggering event of reactivation is not known. Hahn and colleagues20
demonstrated in vitro reactivation of latently infected
granulocyte-macrophage progenitors in the presence of interferon- We have also demonstrated by viral DNA transcription and in vitro infectivity that the increase in DNAemia can produce competent virus in some CMV seropositive individuals. We have not investigated these effects in serologically negative subjects, and do not know if there could be a similar phenomenon in such a cohort. However, others have reported detectable CMV DNA in seronegative individuals: Larsson 19 of 140 (13.6%)26 and Taylor-Weideman 3 of 9 (33%).27 Reactivation with transient DNAemia associated with competent virus may suggest a variation in infectivity risk for blood products from both seropositive and seronegative blood donors. Antibody response to a reactivation event was not reported in these studies nor was it determined in the present work. If development of antibody is delayed in response to reactivation events, a window period of potential infectivity could thus be present before a detectable rise in antibody occurs. Therefore, LR may also add another level of protection for blood products from seronegative donors. This suggests an area for further investigation. Although we have shown that some of the subjects in this study with high CMV levels had competent virus in vitro, we cannot generalize this observation. Further, we have not shown that the resultant levels present after LR are infective or not infective in vivo. To our knowledge, a minimal in vivo infective dose for CMV has not been determined. Our own in vitro data suggest high levels are required before any competency can be demonstrated. One sample at 4200 GE/mL with detectable IE-mRNA did not culture out after 10 days nor did the mRNA persist. The residual CMV levels for the 3 LR methods (apheresis, filtered platelets, and filtered red cells) were pooled and evaluated as a log-normal distribution. This analysis suggests more than 99.9% of leukocyte-reduced products would be expected to be less than 500 GE/mL. This substantial reduction of viral load in leukocyte-reduced blood products supports the clinical efficacy in avoidance of CMV transmission observed in several studies, reviewed by Preiksaitis.28 However, the data presented in this study are not conclusive evidence that LR provides absolute protection against CMV infection for reasons mentioned above: sample size constraints on the estimated proportion of products with detectable CMV DNA (Table 3) and unknown minimal in vivo infective dose. One potential source of bias in this study was that the laboratory performing QA-PCR was not fully blinded to the source or identification of the study samples. We do not feel this is a significant risk to the results, however, because of the objective nature of the measurements; negative and positive controls were run with each batch as described previously, and samples were encoded and processed by different individuals within the laboratory. The primary objective of this study was to test the difference in the effectiveness of 3 LR methods in reducing CMV levels. These methods are used widely for preparation of blood products for indications such as avoidance of nonhemolytic febrile transfusion reactions, alloimmunization, and immunomodulation. Several groups have attempted to subtype the residual WBCs in these blood products and have suggested that there are substantial differences between preparation methods based on their observations. This line of thought has been extended to suggest that these differences are great enough to result in significant differences in clinical outcomes. Residual WBC in leukocyte-reduced blood products are rare events and present substantial challenges in simply enumerating the WBC. The next level of subtyping offers even greater technical challenges. This is evidenced by the lack of repeatability between groups that have attempted this arduous task. For example, in Amicus apheresis products, Tiulzi et al29 found 5 granulocytes/mL, 31 monocytes/mL, and 217 B cells/mL, whereas Johnson and colleagues30 observed 21 granulocytes/mL, 2 monocytes/mL, and 873 B cells/mL. For CMV transmission, this trail of logic requires further assumptions regarding the location and activity of latent virus. Although substantial progress has been made over the past few years in localizing predominant cell types harboring the latent virion31 and the configuration of the genome,32 whether these cell types are the only site of latency or competent virus is still controversial. This study attempted to avoid issues of rare WBC event isolation and subtyping as well as assumptions regarding viral vector location. Instead, we looked specifically for CMV genome in the aggregate final product. Quantitative molecular biology methods were enhanced by extracting nucleic acids from large volumes of final products, with a resulting lower limit of detection of 4 GE/mL. We found a substantial reduction in viral genome load during periods of reactivation in the donor, with more than 99.9% of leukocyte-reduced products expected to have less than 500 GE/mL of CMV DNA. Further, we found apheresis platelets, filtered platelets prepared from whole blood, and filtered RBCs prepared from whole blood to be equivalent in residual CMV DNA in the final product.
The authors would like to thank the apheresis and research staffs of Bonfils Memorial Blood Center and the American Red Cross, Mid-Atlantic Region for their contribution to this work. Carol Afflerbach, Deanna McNeil, Sherrie Sawyer, Bonnie Zishka, and Yvonne Runyan made special contributions that made this logistically difficult study possible. Renee Louis performed the cytokine assays, and Art Hamstra performed WBC counting. Robert Schuyler made very helpful suggestions on the final manuscript.
Submitted July 6, 2000; accepted January 22, 2001.
L.J.D. and T.V.B. are employed by Gambro BCT.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Larry J. Dumont, 10811 Collins Ave, Lakewood, CO 80215; e-mail: larry.dumont{at}gambrobct.com.
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  M. R. Visconti, J. Pennington, S. F. Garner, J.-P. Allain, and L. M. Williamson Assessment of removal of human cytomegalovirus from blood components by leukocyte depletion filters using real-time quantitative PCR Blood, February 1, 2004; 103(3): 1137 - 1139. [Abstract] [Full Text] [PDF]
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W. G. Nichols, T. Price, M. Boeckh, R. Vij, J. DiPersio, H. Khoury, L. T. Goodnough, S. M. Devine, W. Blum, and D. Adkins Donor serostatus and CMV infection and disease among recipients of prophylactic granulocyte transfusions Blood, June 15, 2003; 101(12): 5091 - 5092. [Full Text] [PDF]
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W. G. Nichols, T. H. Price, T. Gooley, L. Corey, and M. Boeckh Transfusion-transmitted cytomegalovirus infection after receipt of leukoreduced blood products Blood, May 15, 2003; 101(10): 4195 - 4200. [Abstract] [Full Text] [PDF]
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B. Wenz, G. A. Ortolano, J.-P. Allain, and L. M. Williamson Leukocyte reduction and HTLV-I: is the glass half empty or half full? Blood, January 1, 2003; 101(1): 370 - 370. [Full Text] [PDF]
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Top
Abstract
Introduction
) and 2 platelet experiments (
). Signal was not detectable
above a background of 200 GE, thus the limit of detection is 4 GE/mL.
minimum detection level of 100 GE/mL). Twenty Norfolk CMV
seropositive subjects were entered into the monitoring phase March 2 to
30 and followed for a maximum of 65 days, through May 5. Twenty-five
Denver CMV seropositive subjects were entered into the monitoring phase
March 15 to April 23 and followed for a maximum of 120 days, through
July 7. Norfolk subjects reactivated during April and Denver subjects
during June/July. Denver had several short-lived increases in tree
pollen throughout the spring, but the largest, sustained period
immediately preceded the subject CMV titer rise. Key: Norfolk subjects,
(
); Norfolk tree pollen, (
or
TNF-