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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 91 No. 1 (January 1), 1998:
pp. 347-352
Impaired Induction of the CD28-Responsive Complex in
Granulocyte Colony-Stimulating Factor Mobilized CD4 T Cells
By
Junji Tanaka,
Marco Mielcarek, and
Beverly Torok-Storb
From the Clinical Research Division, Fred Hutchinson Cancer Research
Center, Seattle, WA and the Department of Medicine, University of
Washington, Seattle, WA.
 |
ABSTRACT |
Use of the CD28/B7 costimulatory signal for T-cell activation was
analyzed in granulocyte colony-stimulating factor (G-CSF) mobilized
peripheral blood mononuclear cells (G-PBMCs) and in peripheral blood
mononuclear cells obtained before administration of G-CSF (preG-PBMCs).
CTLA4Ig inhibition of OKT3-stimulated proliferation was significantly
lower in G-PBMCs compared with preG-PBMCs (39.9% ± 5.6% and 72.2%
± 5.4%, respectively; P < .001). Furthermore, as shown in
electrophoretic mobility-shift assays, the inducible level of the
T-cell transcription factor CD28 responsive complex (CD28RC) was
suppressed in CD4 cells derived from G-PBMC. However, depletion of CD14
cells from G-PBMCs restored CD28RC induction to normal levels. Taken
together, these findings suggest that the large number of CD14
monocytes in G-PBMCs may limit T-cell responsiveness by suppressing the
induction of the CD28RC.
| |
INTRODUCTION |
GRANULOCYTE colony-stimulating factor
(G-CSF) mobilized peripheral blood mononuclear cells (G-PBMCs) have
been used increasingly to reconstitute hematopoiesis after
myeloablative therapy. Although G-PBMC grafts contain at least 10 times
more T cells than standard marrow grafts, the incidence and
severity of acute graft-versus-host disease (aGVHD) is not higher than
observed with allogeneic marrow.1-7 Hypothetically these
clinical observations could be explained by a direct effect of G-CSF on
T-cell function.8 Alternatively, G-PBMCs may contain cells
that can suppress donor T-cell responsiveness.
Previously we have reported that G-PBMC leukapheresis products contain
a large number of CD14+ monocytes that suppress donor
T-cell proliferation in a dose-dependent fashion.9-11
Normal CD14 cells when used in comparable numbers could also suppress
T-cell responsiveness, suggesting that G-PBMCs differed from normal
marrow or PBMCs in the number of CD14 cells. However, our data also
suggested that G-PBMC-derived CD14 cells expressed significantly lower
levels of the costimulatory molecule B7-2 (CD86).9
Given that the engagement of the T-cell receptor in vitro in the
absence of costimulatory molecules can result in a state of
antigen-specific anergy,12-21 we hypothesized that the
large number of CD14+/B7-2lo cells in G-PBMCs
may contribute to low T-cell responsiveness by providing suboptimal
amounts of costimulatory signals. The potential relevance of
costimulatory signals for the activation of donor T cells and the
development of GVHD has been shown in vivo by blocking the interaction
of B7-1 and B7-2 with CD28 on T cells using CTLA4Ig, a soluble fusion
protein of human CTLA-4 and IgG1 Fc region.22 This
treatment reduced lethal aGVHD after allogeneic marrow transplantation
in mice.23,24
Recently, a CD28-responsive element (CD28RE) with promoter activity in
the interleukin-2 (IL-2) gene has been identified17 and
nuclear transcription factors that bind to CD28RE were found and
classified as members of the nuclear factor (NF)  B
family (CD28 responsive complex [CD28RC]).25,26 In the
present study, we have analyzed the role of CD28/B7 costimulation in
T-cell activation by measuring the inducible levels of CD28RC. We show
in CTLA4Ig inhibition studies that CD4 cells in G-PBMCs (G-CD4 cells)
use the CD28/B7 costimulatory pathway to a lesser degree compared with
CD4 cells in preG-PBMCs (preG-CD4 cells). Further, the induction of
CD28RC in G-CD4 cells appears to be suppressed by the presence of CD14
cells in the G-PBMCs.
| |
MATERIALS AND METHODS |
Donors, G-CSF mobilization, and PBMC processing.
Samples were collected from normal volunteers or peripheral blood stem
cell donors after written informed consent was obtained as approved by
the Institutional Review Board of the Fred Hutchinson Cancer Research
Center (Seattle, WA). Heparinized peripheral blood samples were
obtained before the first administration of G-CSF (preG-PBMC).
Peripheral blood stem cell donors were administered recombinant human
G-CSF (rhG-CSF; Amgen, Inc, Thousand Oaks, CA) by
subcutaneous injection at a dose of 8 µg/kg twice daily for 4 to 7
days. Leukapheresis was performed using a continuous flow blood cell
separator (Cobe Laboratories, Lakewood, CO) on 2 consecutive days
beginning on day 4 of rhG-CSF administration, and G-PBMCs were obtained
from the first leukapheresis. PreG-PBMCs were isolated over Ficoll
(Accu-Prep; Accurate Chemicals, Westbury, NY; 1.077-g/mL gradients) and
washed with Hanks' balanced salt solution (HBSS)/1% bovine serum
albumin (BSA). G-PBMCs were suspended in HBSS/1% BSA and centrifuged
at 200 g for 10 minutes to remove platelets and then hemolysed in
hemolysis buffer (150 mmol/L ammonium chloride; 12 mmol/L sodium
bicarbonate). All samples were cryopreserved to allow simultaneous
testing, and paired preG- and G-PBMC samples from the same donor were
used in some experiments as indicated.
Mixed lymphocyte culture (MLC).
Responder PBMCs (25 to 100 × 103) were cultured with
100 × 103 irradiated (30 Gy), allogeneic PBMC
stimulators in 200 µL of RPMI 1640 supplemented with 10% fetal calf
serum (FCS), 0.4 µg/mL L-glutamine, 100 U/mL penicillin, and 100
µg/mL streptomycin in round-bottom 96-well plates (Corning, New
York). After 5 days incubation at 37°C in 5% CO2,
cultures were pulsed with 1.0 µCi/well 3H-thymidine for
the final 16 hours. Cells were harvested and 3H-thymidine
incorporation was measured by scintillation counting. For CTLA4Ig
(kindly provided by Bristol-Myers Squibb, Princeton, NJ)19
inhibition of T-cell proliferation in MLC, both responder and
irradiated stimulator cells were preincubated with 10 µg/mL of
CTLA4Ig for 30 minutes at 37°C.
Polyclonal T-cell stimulation assays using immobilized OKT3
monoclonal antibody.
Ninety-six-well flat-bottomed microtiter plates (Costar, Cambridge,
MA) were preincubated with 6.25 to 100 ng/mL OKT3
monoclonal antibody (Ortho, Raritan, NJ)27 in 100 mmol/L
Tris-HCl buffer (pH 9.5) for 16 hours at 4°C. For CTLA4Ig
inhibition of T-cell responsiveness, cells were preincubated with 0.1
to 30 µg/mL CTLA4Ig for 30 minutes at 37°C before seeding into
OKT3-coated wells. PreG-PBMCs and G-PBMCs were suspended in RPMI 1640
medium supplemented with 10% FCS at indicated concentrations, cultured
at 37°C for 3 days, and pulsed with 3H-thymidine for
the final 16 hours of culture.
Percent CTLA4Ig-mediated suppression was calculated using the following
formula: [3H-thymidine uptake after OKT3 stimulation
without CTLA4Ig (cpm)  OKT3 stimulation with CTLA4Ig
(cpm)]/OKT3 stimulation without CTLA4Ig (cpm) × 100.
To analyze induction of nuclear transcription factor CD28RC in
preG-PBMCs and G-PBMCs, 1 × 106/mL cells were
cultured in OKT3-coated (25 ng/mL) T75 culture flasks (Costar) for 4
hours with or without CTLA4Ig (10 µg/mL).
Immunofluorescent staining and flow cytometric analysis.
Cells were stained with Leu-3a (anti-CD4-phycoerythrin
[PE]; Becton Dickinson, San Jose, CA) plus Leu-28
(anti-CD28-fluorescein isothiocyanate [FITC], Becton Dickinson) and
with LeuM3 (anti-CD14-PE, Becton Dickinson) plus MAB104 (anti-CD80
[B7/BB1]-FITC, Immunotech, Boston, MA) or FUN-1 (anti-CD86
[B70/B7-2]-FITC, Pharmingen, San Diego, CA). Isotype-matched
antibodies of irrelevant specificity were used as negative controls.
Samples were analyzed with the use of a FACS Calibur (Becton Dickinson)
flow cytometer.
Immunomagnetic cell sorting for CD4 enrichment and CD14 depletion.
Enriched CD4 T cells were obtained by positive selection using Leu-3a
anti-CD4-FITC, antihuman-mouse IgG1 (Becton Dickinson) as primary
antibody and rat-antimouse IgG1 conjugated to magnetic microbeads as
the secondary antibody. Flow cytofluorometric analysis of enriched
populations indicated that they contained >90% CD4+
cells. CD14 cell-depleted fractions containing <1% CD14+
cells were obtained by negative selection using LeuM3 anti-CD14-PE,
antihuman-mouse IgG2a (Becton Dickinson) as primary antibody and
rat-antimouse IgG2a+b conjugated to magnetic microbeads as the
secondary antibody according to the manufacturer's instructions
(Miltenyi Biotec GmbH, Bergisch Gladbach, Germany).
Electrophoretic mobility-shift assay (EMSA).
Unfractionated preG- or G-PBMCs were cultured for 4 hours on
immobilized OKT3 antibody. CD4 cells were then purified by
immunomagnetic enrichment as described previously, and nuclear extracts
were isolated from purified CD4 cells as described
previously.28,29 In brief, cells were suspended in 400 µL
of cold hypotonic buffer containing 10 mmol/L HEPES-KOH pH 7.9, 10
mmol/L KCI, 0.1 mmol/L EDTA, 0.1 mmol/L EGTA, 1 mmol/L dithiothreitol
(DTT), and 0.5 mmol/L phenylmethylsulfonyl fluoride (PMSF). The cells
were allowed to swell on ice for 20 minutes. Thereafter, 25 µL of
10% NonidentP-40 (Sigma, St Louis, MO) was added and the contents were
vigorously mixed. After centrifugation, the nuclear pellet was
resuspended in 50 µL of cold extraction buffer (20 mmol/L HEPES-KOH,
pH 7.9, 400 mmol/L NaCl, 1 mmol/L EDTA, 1 mmol/L EGTA, 1 mmol/L DTT,
and 1 mmol/L PMSF) and vigorously rocked at 4°C for 60 minutes.
Nuclear extracts were cleared by centrifugation. Protein concentration
was assayed using a bicinchoninic acid (BCA) protein assay kit (Pierce,
Rockford, IL). The oligonucleotide
5 -AAAGAAATTCCAAAGAAAAGAAATTCCAAAGA-328 was
used as a CD28RC recognition sequence after 32P-end
labeling by T4 polynucleotide kinase (GIBCO-BRL, Gaithersburg, MD).
Each reaction mixture contained about 100 fmol of double-stranded
32P-end labeled probe and 1 µg of protein from nuclear
extracts. Binding reactions were performed in binding buffer (25 mmol/L
HEPES-KOH, pH 7.9, 30% [vol/vol] Glycerol, 10 mmol/L
MgCl2, 50 mmol/L KCl, 1 mmol/L EDTA, 1 mmol/L DTT, and 0.5
µg of poly[dI-dC]) in a final volume of 20 µL at room temperature
for 20 minutes. Complexes of nuclear protein and 32P-end
labeled probe were electrophoresed on 6% polyacrylamide gels in
TBE buffer (22 mmol/L Tris, 22 mmol/L borate, and 0.4
mmol/L EDTA) for about 2 hours (100 constant voltage [CV]). Gels were
dried and analyzed using ImageQuant3.3 software (Molecular Dynamics,
Sunnyvale, CA).
| |
RESULTS |
Proliferative responsiveness of preG- and G-PBMCs in MLC.
Unfractionated G-PBMC showed lower proliferative responses in MLC
compared with equivalent numbers of unfractionated preG-PBMCs obtained
from the same donor (Fig 1).
Hyporesponsiveness of G-PBMCs in MLC may be caused by the lower
percentage of T cells and the higher percentage of suppressive CD14
cells in G-PBMCs compared with preG-PBMCs, as reported
previously.9 In addition, CTLA4Ig-mediated inhibition in
MLC was more pronounced at all responder cell concentrations when
preG-PBMCs compared with G-PBMCs were used as responders.
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| Fig 1.
Mixed lymphocyte cultures using increasing numbers of
preG-PBMCs (A) and G-PBMCs (B) cultured with 100 × 103
irradiated, allogeneic PBMC stimulators in the presence (solid line) or
absence (dashed lines) of 10 µg/mL CTLA4Ig. Values for
3H-thymidine incorporation represent the mean counts per
minute (cpm) ± the standard error of the mean (SEM) from triplicate
cultures. Results from one of three representative experiments are
shown.
|
|
PreG- and G-PBMC proliferation in response to immobilized OKT3
monoclonal antibody.
Unfractionated preG- and G-PBMCs obtained from the same donor were
stimulated with immobilized OKT3 monoclonal antibody at different
concentrations (Fig 2
[View Larger Version of this Image (11K GIF file)]
A and
B). OKT3 monoclonal antibody immobilized at a concentration of 25 ng/mL
provided suboptimal stimulation, which could be largely inhibited by 10
µg/mL CTLA4Ig in preG-PBMCs. CTLA4Ig-mediated inhibition was less
effective at higher OKT3 concentrations. Therefore, an OKT3
concentration of 25 ng/mL was chosen for subsequent polyclonal
stimulation assays.
Polyclonal T-cell stimulation assays using
immobilized OKT3 monoclonal antibody. Unfractionated preG-PBMCs (A) and
G-PBMCs (B) (100 × 103 cells/200 µL) were cultured in
the presence (solid line) or absence (dashed line) of 10 µg/mL
CTLA4Ig. Values for 3H-thymidine incorporation represent
the mean cpm ± SEM from triplicate cultures. (C) Dose-dependent
CTLA4Ig-mediated inhibition of immobilized OKT3 stimulated preG-PBMCs
(solid line) and G-PBMCs (dashed line) obtained from the same donor.
Cells were stimulated by immobilized OKT3 (25 ng/mL) in the absence or
presence of CTLA4Ig at the concentrations indicated. The percent
CTLA4Ig-mediated inhibition was calculated as described in Materials
and Methods. Results from one of three representative experiments are
shown.
Fig 2.
CTLA4Ig-mediated inhibition of immobilized OKT3-stimulated preG- and
G-PBMC proliferation.
The role of CD28/B7 costimulation in preG- and G-PBMCs was compared by
measuring T-cell proliferation in response to OKT3 stimulation in the
presence and absence of different concentrations of CTLA4Ig. Comparing
paired samples from the same donor, CTLA4Ig consistently had a greater
inhibitory effect on the preG- compared with the G-PBMC proliferative
response. As shown in Fig 2C, maximum inhibition of preG-PBMC
proliferative response was >70% at a CTLA4Ig concentration of 3
µg/mL. In contrast, maximum inhibition of G-PBMC proliferation was
<30% even at a CTLA4Ig concentration of 30 µg/mL.
In larger series of experiments, CTLA4Ig at 10 µg/mL was used to
evaluate the role of B7 costimulation in OKT3-stimulated
proliferation in both paired and unpaired samples of preG- and G-PBMCs.
Using all samples the inhibition of proliferation by CTLA4Ig in
preG-PBMCs (n = 11) was 72.2% ± 5.4%, which was significantly
higher than the 39.9% ± 5.6% (P < .001) observed in
G-PBMCs (n = 15). When the five paired samples were analyzed
separately, CTLA4Ig inhibition in preG-PBMCs was also significantly
higher than that seen in G-PBMCs (62.9% ± 10.0% v 25.4%
± 11.6%; P < .002; data not shown).
Expression of costimulatory molecules in preG- and G-PBMCs.
Expression levels for CD28/B7 were compared between CD4 and CD14 cells
derived from preG- and G-PBMCs. As reported previously,9
CD28 was equally expressed on preG- and G-CD4 cells. In addition, preG-
and G-CD14 cells had comparably low expression of B7-1 (CD80). However,
expression levels of B7-2 (CD86), a ligand for CD28, were reduced by
60% to 70% on G-CD14 cells compared with preG-CD14 cells (data not
shown).
Detection of CD28RC in EMSA.
CD28RC was detectable in the nuclear protein fraction extracted from
CD4 T cells derived from OKT3-stimulated preG-PBMCs by this assay (Fig
3). The addition of unlabeled CD28RE
inhibited the detection of CD28RC in a dose-dependent manner. However,
the addition of a nonspecific probe did not interfere with the
detection of CD28RC, indicating that this assay was specific for
CD28RC.
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| Fig 3.
Detection of CD28RC in EMSA. The nuclear extract from CD4
cells derived from OKT3-stimulated preG-PBMCs was incubated with a
32P-labeled CD28RE. Extracts were electrophoresed on a 6%
polyacrylamide gel and complexes of nuclear proteins and CD28RE were
visualized using ImageQuant3.3 software (Molecular Dynamics). Unlabeled
CD28RE (10 and 50 pmol) as a specific competitor or negative regulatory
element A (NRE-A) as a nonspecific competitor were added as
indicated.
|
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Comparison of CD28RC induction in CD4 T cells derived from
OKT3-stimulated preG- and G-PBMCs.
Because of functional differences seen in the CTLA4Ig inhibition
experiments, we analyzed CD28RC expression in CD4 cells derived from
unfractionated preG- and G-PBMCs after 4 hours of stimulation on
immobilized OKT3 monoclonal antibody. The purity of CD4 cells after
immunomagnetic enrichment and before nuclear protein extraction usually
exceeded 90% as assessed by flow cytometric analysis. In preG-CD4
cells, CD28RC expression was upregulated by OKT3 stimulation and
suppressed in the presence of CTLA4Ig (Fig
4). In contrast, CD28RC expression in G-CD4
cells was not significantly increased in responses to OKT3 stimulation.
In addition, the inhibitory effect of CTLA4Ig on CD28RC expression was
considerably less pronounced in G-CD4 compared with preG-CD4 cells.
Thus, the inducible expression of CD28RC appeared to be suppressed in
G-CD4 cells.
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| Fig 4.
EMSA of CD28RC in OKT3-stimulated preG- and G-CD4 cells.
1 × 106/mL unfractionated preG-PBMCs and G-PBMCs were
cultured for 4 hours in OKT3-coated T75 culture flasks in the presence
or absence of CTLA4Ig. One microgram of nuclear extract isolated from
CD4 cells purified by immunomagnetic enrichment was used for the EMSA.
Results from one of four experiments are shown.
|
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Proliferation of CD14-depleted G-PBMCs in response to stimulation
with immobilized OKT3.
We have reported previously that G-PBMCs contain a large number of
CD14+ monocytes that suppress T-cell proliferation in MLC
in a dose-dependent manner.9 To analyze whether CD14 cells
can also suppress OKT3-stimulated T-cell proliferation, we compared
unfractionated and CD14-depleted G-PBMCs. As shown in Fig
5, CD14-depleted G-PBMCs (<1% CD14
cells) responded better than unfractionated G-PBMCs to OKT3
stimulation. In addition to the improved responsiveness, CD14-depleted
G-PBMCs were more susceptible to CTLA4Ig-mediated inhibition. In
contrast, OKT3-stimulated proliferation of unfractionated G-PBMCs
decreased at cell concentrations exceeding 100 ×
103/200 µL.
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| Fig 5.
3H-thymidine incorporation by OKT3-stimulated
G-PBMCs. Unfractionated G-PBMCs (A) and CD14 depleted G-PBMCs (B) were
cultured at various cell concentrations in the absence (solid lines) or
in the presence (dashed lines) of CTLA4Ig. Values for
3H-thymidine incorporation represent the mean cpm ± SEM
from triplicate cultures. Results from one of four representative
experiments are shown.
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CD28RC induction in G-CD4 cells derived from unfractionated and CD14
depleted G-PBMCs in response to stimulation with immobilized OKT3.
To test whether CD14+ monocytes in G-PBMCs could suppress
OKT3-induced expression of CD28RC, CD4 cells derived from
unfractionated and CD14-depleted G-PBMCs were analyzed for CD28RC
expression. As shown in Fig 6, this
transcription factor was expressed at considerably higher levels in CD4
cells derived from CD14-depleted compared with unfractionated G-PBMCs.
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| Fig 6.
EMSA of CD28RC in OKT3-stimulated CD4 cells derived from
unfractionated or CD14-depleted G-PBMCs. A total of 1 ×
106/mL unfractionated G-PBMCs and CD14 depleted G-PBMCs
were cultured in OKT3-coated T75 culture flasks for 4 hours. One
microgram of nuclear extract isolated from CD4 cells derived from
either unfractionated (lane 1) or CD14-depleted (lane 2) G-PBMCs was
used for EMSA. Results shown are generated from the same sample used in
the experiment shown in Fig 5.
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| |
DISCUSSION |
The present study investigates the role of the CD28/B7-costimulatory
pathway in T-cell activation in G-PBMCs compared with normal peripheral
blood. This study was prompted by the observations that unfractionated
G-PBMCs are hyporesponsive to alloantigen stimulation in vitro and that
the monocytes that comprise 25% to 50% of the cells in G-PBMCs
express reduced levels of B7-2 (CD86), a critical costimulatory
molecule for T-cell activation.9 Using OKT3 stimulation and
CTLA4Ig inhibition experiments, we now report that in comparison to
normal PBMCs, G-PBMCs are less susceptible to CTLA4Ig-mediated
suppression of OKT3-stimulated proliferation. This suggests that
CD28/B7 interactions may not contribute significantly to T-cell
activation in G-PBMCs. In keeping with this observation, G-CD4 cells
have decreased levels of inducible CD28RC. However, the inducible
levels of this DNA binding protein increased after G-PBMCs were
depleted of CD14 cells. These data suggest that the large number of
CD14+/B7-2lo cells in G-PBMCs may be
interfering with CD28 signal transduction.
B7-1 and B7-2 molecules expressed on antigen-presenting cells (APCs)
bind to CD28 and CTLA-4 on T cells, thereby providing a costimulus for
T-cell activation and clonal expansion.12-14,17 Antigen
presentation by APCs to T cells without costimulation results in a
state of antigen-specific anergy. It has been shown in vitro that
blocking the interaction between CD28 and B7 can result in T-cell
clonal anergy.18-21 In addition, in vivo data suggest that
blockade of the CD28/B7 costimulatory pathway with CTLA4Ig, which
specifically blocks B7-1 and B7-2 on the APC side, may reduce lethal
aGVHD after allogeneic marrow transplantation in mice.23,24
Therefore, costimulatory molecules may play an important role in
regulating T-cell function and the development of GVHD in human marrow
and blood stem cell transplantation.30 The unexpectedly low
incidence and severity of aGVHD in patients receiving G-PBMCs despite
the transfer of at least 10 times more T cells compared with marrow
transplantation may be a consequence of the large number of
CD14+/B7-2lo cells suppressing CD28-mediated
signal transduction.
Whether the relatively low levels of B7-2 expressed by these cells
contributes directly to this suppressive effect is not clear at this
time. Monocytes, in addition to their role as APCs,31,32
can also inhibit T-cell function by several
mechanisms.33-36 Soluble monocyte products have been shown
to downregulate nuclear transcription factors required for T-cell
activation and proliferation. IL-10, for example, inhibits NF-B/Rel
activity and prostaglandin E2 inhibits AP-1 and NF-AT induction in
human T cells.37,38 Therefore, in addition to the potential
relevance of reduced B7-2 levels, factors produced and secreted by
monocytes may play a role in downregulation of CD28RC in G-CD4 cells.
There also remains the possibility that G-CSF treatment might directly
alter T-cell function by altering signaling events downstream of the
CD28 receptor.
These considerations notwithstanding, our data suggest a decreased use
of the CD28/B7 costimulatory pathway by stimulated CD4 T cells in
G-PBMC leukapheresis products which, in turn, appears to be influenced
by the large number of CD14/B7-2lo cells present. Whether
these findings influence either the incidence or severity of aGVHD
observed after peripheral blood stem cell transplantation remains
speculative. However, they may stimulate interest in investigating the
potential use of manipulated cell populations to prevent or treat
graft-versus-host reactions.
| |
FOOTNOTES |
Submitted June 16, 1997;
accepted September 2, 1997.
Supported in part by grants DK51417 and CA18221 awarded by the National
Institutes of Health, DHHS, Bethesda, MD. J.T. is supported by the
Ministry of Education, Science and Culture, Tokyo, Japan.
Address reprint requests to Junji Tanaka, MD, PhD, Fred Hutchinson
Cancer Research Center, 1124 Columbia Street, M318, Seattle, WA 98104.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. section 1734 solely
to indicate this fact.
| |
ACKNOWLEDGMENT |
We thank Gretchen Johnson, Ludmila Golubev, and Laura Bolles for their
technical assistance and Harriet Childs for typing the manuscript.
| |
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Abstract
Introduction
Abstract