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Blood, Vol. 96 No. 2 (July 15), 2000:
pp. 754-762
TRANSPLANTATION
Prolonged CD4 depletion after sequential autologous peripheral
blood progenitor cell infusions in children and young adults
Crystal L. Mackall,
Dagmar Stein,
Thomas A. Fleisher,
Margaret R. Brown,
Frances T. Hakim,
Catherine V. Bare,
Susan F. Leitman,
Elizabeth J. Read,
Charles S. Carter,
Leonard H. Wexler, and
Ronald E. Gress
From the Pediatric Oncology Branch, Medicine Branch, Experimental
Immunology Branch, National Cancer Institute and Department of
Transfusion Medicine, Clinical Center, National Institutes of Health,
Bethesda, MD.
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Abstract |
Administration of mobilized peripheral blood progenitor cells
(PBPCs) after high-dose chemotherapy rapidly restores multilineage hematopoiesis, but the ability of such products to restore lymphocyte populations remains unclear. In this report, we evaluated immune reconstitution in a series of patients treated with sequential cycles
of high-dose chemotherapy, followed by autologous PBPC infusions
(median CD34+ cell dose
7.2 × 106 cells/kg [range 2-29.3]).
Although patients experienced rapid reconstitution of B cells and
CD8+ T cells, we observed CD4 depletion and diminished
immune responsiveness in all patients for several months after
completion of therapy. Mature CD4+ T cells contained
within the grafts did not appear to contribute substantially to immune
reconstitution because CD4 counts did not differ between recipients of
unmanipulated T-cell replete infusions versus CD34 selected,
T-cell-depleted infusions. Rather, at 12 months after therapy, total
CD4 count was inversely proportional to age ( =  0.78,
P = .04), but showed no relationship to CD34 cell dose
( = 0.42, P = .26), suggesting that age-related
changes within the host are largely responsible for the limited immune reconstitution observed. These results demonstrate that in the autologous setting, the infusion of large numbers of PBPCs is not
sufficient to restore T-cell immune competence and emphasize that
specific approaches to enhance immune reconstitution are necessary if
immune-based therapy is to be used to eradicate minimal residual
disease after autologous PBPC transplantation.
(Blood. 2000;96:754-762)
© 2000 by The American Society of Hematology.
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Introduction |
There are 2 primary pathways by which CD4+
T cells may be regenerated in vivo. Thymic-dependent CD4
regeneration recapitulates immune ontogeny and gives rise to peripheral
T cells via thymic differentiation of primitive marrow-derived
progenitors. Alternatively, peripheral expansion of mature
CD4+ T cells can also restore substantial CD4+
T-cell numbers post-T-cell depletion, and this represents the primary
pathway for CD4+ T-cell regeneration when thymopoietic
pathways are limiting.1
We and others have previously shown that persistent CD4 depletion after
intensive chemotherapy and after bone marrow transplantation (BMT) in
humans is related primarily to age-associated thymic insufficiency.2-4 The mechanisms responsible for
age-associated thymic insufficiency are poorly understood, but could
involve age-associated declines in bone marrow-derived T-cell
progenitors that may be intensified by cytotoxic therapy. Recent
evidence has shown that in the allogeneic transplant setting, the
provision of mobilized peripheral blood progenitor cells (PBPCs) leads
to more rapid immune reconstitution compared with BMT5,6
and some reports have suggested similar improvements in immune recovery in autologous PBPC transplantation compared with autologous
BMT.7,8 Such enhanced T-cell regeneration could result from
the provision of larger numbers of prethymic progenitors, which could
enhance the regeneration of mature T cells via thymic pathways.
Alternatively, however, enhanced immune reconstitution after PBPC
transplantation versus BMT could result from the provision of larger
numbers of T cells contained within PBPC grafts compared with bone
marrow grafts,9 which might serve to enhance T-cell
reconstitution via peripheral expansion.
In this study, immune reconstitution was evaluated in a series of
children and young adults treated for cancer with sequential cycles
of dose-intensive chemotherapy, followed by mobilized autologous PBPC
infusions. The goals of the study were to investigate whether the
provision of PBPCs was sufficient to restore T-cell populations that
were depleted by high-dose alkylating agent therapy and to address
whether such PBPC populations abrogated the age-associated declines in
thymic regenerative capacity, which we have reported previously.
Furthermore, because mature T cells contained within PBPC grafts could
contribute to immune reconstitution with resultant important
implications for host immune competence, we sought to address whether
immune reconstitution differed between recipients of T-cell-replete
versus T-cell-depleted PBPC grafts.
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Patients, materials, and methods |
Patients and chemotherapy
The patient population comprised children and young adults treated
for pediatric sarcomas (Ewing sarcoma family of tumors, rhabdomyosarcoma) or neuroblastoma. The patients were treated on
protocol NCI 93-C-0125, which was approved by the Institutional Review
Board of the National Cancer Institute and informed consent was
obtained from all patients or their parents before enrollment. The
trial was open to patients aged 1 to 25 years. All patients enrolled in
this trial of high-dose therapy were at high risk for tumor recurrence
either because of age (more than 10 years), tumor location (truncal or
proximal extremity), or disease extent (regional or disseminated). No
patient received chemotherapy before enrollment on this protocol.
Patients received chemotherapy according to 2 phases: an
induction/progenitor cell mobilization phase (cycles 1-5) during which
dose-intensive, cytotoxic therapy was administered and PBPCs were
collected by apheresis, followed by a consolidation phase (cycles 6-8)
during which 1 to 3 cycles of high-dose chemotherapy, followed by PBPC
infusion were administered. The number of consolidation cycles
administered to each patient was based on patient tolerance and
CD34+ cell availability. Induction therapy comprised
vincristine 2.0 mg/m2 (max 2.0 mg), administered weekly
× 12 and, simultaneously, 5 consecutive cycles of doxorubicin 90 mg/m2 and cyclophosphamide 2.4 gm/m2 (VadriaC),
with each cycle commencing as soon as possible after hematologic
recovery. Granulocyte colony-stimulating factor (G-CSF) 10 mcg/kg was
administered daily after each cycle through recovery of the absolute
neutrophil count (ANC) to at least 10 000 cells/µL or until
completion of the apheresis procedure for a given cycle. During the
consolidation phase, therapy consisted of up to 3 consecutive cycles of
melphalan (dosage range of 60-120 mg/m2), ifosfamide 13.6 gm/m2, etoposide 800 mg/m2, and mesna as a
uroprotectant (MIME). At least 72 hours after infusion of MIME
chemotherapy, PBPCs were administered, followed by G-CSF 10 mcg/kg per
day. Radiotherapy (5.2-6.0 Gy) was administered to the site of the
primary tumor in patients 1, 6, 7, 11, 13, 14, and 19. Because
pneumonia with Pneumocystis carinii developed in 2 of the first
5 patients treated on this trial, subsequent patients were treated
prophylactically with either inhaled pentamadine (300 mg) or
trimethoprim/sulfamethoxazole (150/750 mg/m2 per day)
administered orally twice weekly.
Collection and processing of autologous peripheral blood
progenitor cells
PBPCs were collected by automated leukapheresis using the Fenwal
CS3000Plus (Baxter, Deerfield, IL) or the Spectra (Cobe, Lakewood, CO)
cell separator by peripheral or central venous access. Citrate
anticoagulation was used, but was occasionally reduced and augmented
with heparin in smaller patients to avoid citrate toxicity. Blood
volume processed was typically 2 to 3 blood volumes (10-15 L for
adults) per apheresis procedure. Leukapheresis was initiated at the
beginning of recovery from the leukocyte nadir after induction
chemotherapy, when the circulating leukocyte count exceeded
2000/µL, and was continued daily to achieve a target CD34+ cell dose of 5 × 106/kg patient
weight. In 1 patient (no 9), adequate numbers of CD34+
cells were not obtained via harvests undertaken during induction chemotherapy, therefore multiple subsequent aphereses were performed after MIME consolidation chemotherapy. Individual patients underwent a
mean of 4.05 ± 0.95 PBPC apheresis procedures with
11.36 ± 1.41 L processed/procedure.
PBPCs were not CD34 selected for the first 9 patients, but were
CD34 selected using the CeprateSC system (CellPro,
Bothell, WA) for the remaining 10 patients. This selection method,
which uses a biotinylated anti-CD34 antibody, purifies cells by
biotin-avidin immunoadsorption and typically results in 50% recovery
of CD34+ cells, and a 2-log passive depletion of T cells.
All unselected and selected PBPC products were cryopreserved in 10%
DMSO using controlled-rate freezing and stored in bags or vials in
liquid nitrogen until ready for thawing and infusion. CD34+
cells and CD3+ T cells in PBPC products were enumerated by
automated leukocyte counting and flow cytometry using the FACScan with
CellQuest software (Becton Dickinson Immunocytometry Systems [BDIS],
San Jose, CA), with standard fluorochrome-labeled antibodies to CD34
(anti-HPCA-2) and CD3 (Leu 4) (BDIS). For each cycle of consolidation
therapy, the CD34+ and CD3+ cell doses for all
bags or vials infused were summed and expressed as a cell dose per
kilogram patient weight (Table 1).
Flow cytometry
Peripheral blood specimens were obtained during routine clinic
visits and were handled according to established clinical guidelines. Baseline samples were obtained before therapy. On-therapy evaluation of
lymphocyte subsets was undertaken at the time of hematologic recovery just before administration of cycles 3, 6, 7, and 8. On
completion of therapy, immunophenotyping was undertaken at the time of
routine clinic visits that generally occurred at 1, 3, 6, 9, and 12 months after therapy, then every 6 months for up to 30 months after
completion of therapy. Evaluation of immune reconstitution was
discontinued at the time of tumor recurrence.
Cells were stained for flow cytometry using the whole blood lysis
technique and analyzed on a FACScan using Cellquest software as
previously described.10 To calculate absolute numbers of each lymphocyte subset, the percentage of cells staining positive was
multiplied by the absolute peripheral blood lymphocyte count. This was
determined by a CellDyn 3500 (Abbott, Chicago, IL) and leukocyte
differential on a blood sample obtained simultaneously. Control samples
obtained from normal volunteers were analyzed concurrently with each
experimental sample. The 95% confidence intervals for lymphocyte
subsets as shown in Figures 1 and
2 were derived from 40 healthy adults aged
18 to 55 in whom lymphocyte subset analysis was performed as described
above.
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| Fig 1.
Changes in peripheral blood lymphocyte subpopulations
during sequential cycles of chemotherapy, PBPC infusions, and after
completion of therapy.
The absolute number of each lymphocyte subset was determined using flow
cytometry as described in "Materials and methods." Samples were
obtained at the time of maximal hematologic recovery from the preceding
cycle of chemotherapy. Open circles represent patients who received
unmanipulated PBPC grafts and filled squares represent patients who
received CD34-selected/T-cell-depleted PBPC grafts. Shaded areas
represent 95% confidence intervals for adult normal values as
described in "Materials and methods." Horizontal lines denote
median values. The time points noted as 1 to 30 months represent time
after completion of PBPC-supported chemotherapy. Patients received 1 to
3 PBPC cycles as detailed in Table 1. Pre-rx n = 15, Cy3 n = 8,
PBPC1 n = 13, PBPC2 n = 8, PBPC3 n = 8, 1 month n = 5, 3 months
n = 12, 6 months n = 5, 9 months n = 4, 12 months n = 8, 15 to
18 months n = 6, 24 months n = 4, and 30 months n = 2.
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| Fig 2.
Changes in peripheral blood T-cell and CD4+
T-cell subsets during sequential cycles of chemotherapy, PBPC
infusions, and after completion of therapy.
The absolute number of each subset was determined using flow cytometry
as described in "Materials and methods." Samples were obtained
at the time of maximal hematologic recovery from the preceding cycle
of chemotherapy. Shaded areas represent 95% confidence intervals for
normal values as described in "Materials and methods." Horizontal
lines denote median values. The time points noted as 1 to 30 months
represent time after completion of PBPC-supported chemotherapy.
Patients received 1 to 3 PBPC cycles as detailed in Table 1. N
values for each time point are shown in the Figure 1 legend.
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The monoclonal antibodies used for T-cell phenotyping included:
anti-CD3(Leu 4), anti-CD4 (13B8.2), anti-CD8 (B9.11), anti-CD45RO (UCHL1), and anti-CD45RA (Alb11). T-cell activation
antibodies included: anti-HLA-DR and anti-CD25 (2A3). Natural killer
(NK) antibodies included: anti-CD16 (Leu 11) and anti-CD56 (Leu 19). B-cell antibodies were anti-CD19 (Leu 12) and anti-CD20 (Leu 19). The
Leu reagents, anti-CD25 and anti-HLA-DR were obtained from Becton
Dickinson; the anti-CD45RO reagent was obtained from Dako, Inc
(Carpinteria, CA), anti-CD45RA was from Beckman Coulter (Brea, CA), and
anti-CD4 and anti-CD8 were from Coulter-Immunotech (Hialeah, FL).
Irrelevant murine monoclonal antibodies of the IgG1, IgG2a, and IgG2b
subclass (BDIS) were used to define background staining.
Lymphocytes were identified by forward- and side-scatter analyses and
the lymphocyte gate was checked using the Leucogate (CD45/CD14) reagent
from Becton Dickinson. List mode parameters were collected for 10 000
cells within the lymphocyte gate and positive staining was calculated
based on the subclass control specimens. CD4+ T cells were
defined as CD4+CD3+ cells, B cells as
CD20+ cells, and NK cells as CD3 cells,
which were CD16+ and/or CD56+. Three-color
analysis was used to define activated T-cell subsets (CD3+/CD4+ or
CD8+/HLA-DR+ or CD25+)
and CD45 isoform expression
(CD3+/CD4+/CD45RA+ or
CD45RO+).
Functional assays
PBPCs from patients and normal donors were separated using
ficoll-sodium diatrizoate gradients (Lymphocyte Separation Medium, Organon Teknika, Durham, NC), washed twice with Delbecco's
phosphate-buffered saline (DPBS) (Gibco/BRL, Gaithersburg, MD), and
resuspended at 2 × 106 cells/mL in RPMI 1640 (Gibco) supplemented with penicillin/streptomycin, and 10% fetal
bovine serum (Sigma, St Louis, MO). Cells were cultured at
2 × 105 cells per well in 96-well round bottom
plates with medium alone and phytohemagglutinin (PHA, 0.5% final,
Gibco). Wells were pulsed after 48 hours with 0.037 Mbq (1 µCi) 3H thymidine and harvested 18 to 24 hours later using a Titertek harvester (Wallac,
Turku, Finland). Thymidine uptake was measured using a
Betaplate reader (Wallac).
Immunoglobulin levels
Serum IgG, IgA, and IgM levels were determined using standard
automated nephelometry at specified time points during chemotherapy and
sequentially on completion of chemotherapy.
Statistics
Data are expressed as medians (ranges). Because analysis of this
data set is limited because of low patient numbers, the nonparametric Spearman correlation coefficients and Mann-Whitney U tests for unpaired comparisons were calculated where described. All P
values are 2-sided.
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Results |
Lymphocyte depletion during induction and consolidation chemotherapy
Patient characteristics are listed in Table 1. This trial was
designed to enroll a minimum of 5 and a maximum of 17 patients of each
of 3 histologies (Ewing sarcoma family of tumors, rhabdomyosarcoma and
neuroblastoma) to evaluate disease response to this regimen of dose
compressed, intensive multimodality therapy. Because significant toxicity was observed without obvious benefit in terms of disease control, the protocol was closed before completing full enrollment. The
19 patients evaluated in this report represent consecutive patients
enrolled for whom immune endpoints were available.
To evaluate the changes in lymphocyte number and phenotype induced by
sequential multiagent chemotherapy, peripheral blood immunophenotyping
was performed at the time of presentation and on hematologic recovery
from successive cycles of induction and consolidation chemotherapy
(Figure 1). At presentation, median B-cell number was 185 cells/µL (range 35-425), NK cell number was 168 cells/µL (range 38-313), CD4+ T-cell number
was 608 cells/µL (range 119-1098) and CD8+
T-cell number was 325 cells/µL (range 90-554). Median
T-cell numbers were within normal adult limits, although moderately
diminished values were recorded in some patients, as previously
reported.10 During the induction phase, patients received 5 cycles of VadriaC therapy, which resulted in profound lymphocyte
depletion in all patients. Depletion was most dramatic in the
T-cell and B-cell subsets with a median CD4+ T-cell count
of 83 cells/µL (range 13-154), CD8+ T-cell count of 104 cells/µL (range 9-857) and a median B-cell count of 8 cells/µL (range 1-101) at the time of hematologic recovery before
the first PBPC-supported cycle of chemotherapy. NK cells were less
severely depleted with a median value of 92 cells/µL (range
9-262) cells at the same time point (Figure 1).
Autologous PBPCs were generally harvested during the recovery phase of
cycles 2, 3, and 4 of VadriaC therapy, at which time significant
changes in T-cell number and phenotype had already taken place. As
shown in Figure 2, there was a dramatic increase in CD25 receptor
expression on T cells after cycle 2 of therapy, indicative of
widespread T-cell activation. Although CD25 receptor expression was
transient, this was followed by a more prolonged expression of HLA-DR
and a conversion to the CD45RO phenotype that persisted for months in
these patients. Therefore, dose-intensive chemotherapy not only induced
dramatic T-cell depletion, but also induced widespread activation of
the T-cell populations remaining after such chemotherapy.
After VAdriaC induction therapy, up to 3 cycles of consolidation
MIME therapy accompanied by peripheral blood progenitor cell infusions were administered. The number of MIME/PBPC cycles given to
each patient and the CD34 dose for each cycle of MIME/PBPC administered
are shown in Table 1. As shown in Table 1, PBPC infusions resulted in
rapid restoration of hematopoiesis with a median duration of
neutropenia (ANC < 500 cells/µL) of 7.0 days (range
5-13), and a median duration to recovery of a transfusion-independent platelet count of more than 50 000 cells/µL
(from day 1 of the cycle) of 16 days (range 2 to more
than 100). Successive cycles of MIME/PBPC were administered upon
hematologic and clinical recovery and occurred at a median of 31 days
(range 23-102). In 2 patients (16 and 17), a second PBPC
infusion was administered at day 43 and day 100, respectively, due
to moderate asymptomatic pancytopenia. In both patients, subnormal
hematologic parameters persisted for several months after the second
PBPC infusion.
Despite rapid restoration of hematopoiesis in the majority of patients
during these sequential consolidation cycles, reconstitution of
lymphocyte populations did not occur. As shown in Figure 1, median
CD4+ T-cell number before PBPC cycle 1 was 83 cells/µL
(range 13-147), whereas median CD4+ T-cell number after
PBPC cycle 1 was 64 cells/µL (range 12-152), after PBPC cycle 2 was
37 cells/µL (range 16-112) and after PBPC cycle 3 was 34 cells/µL
(range 2-145). Similarly, CD20+ B cells remained very low
during the consolidation phase. In contrast, CD8+
T cells and NK cells showed less depletion during MIME/PBPC
consolidation therapy, with median values at 1 month that were only
modestly depleted compared with baseline. Therefore, although PBPC
infusions resulted in rapid restoration of hematopoiesis, allowing
sequential PBPC-supported cycles of dose-intensive chemotherapy to be
administered with relative safety, restoration of CD4+ and
B-cell lymphocyte populations was not observed within the same time frame.
Immune reconstitution after completion of chemotherapy/ peripheral
blood progenitor cell infusions
To characterize the pace and extent of lymphocyte recovery after
autologous PBPC infusions, evaluation of lymphocyte subsets was
performed sequentially on completion of PBPC-supported chemotherapy. However, because tumor recurrence occurred relatively early in several
patients as detailed in Table 1, many patients from the initial cohort
were not available for analysis of late immune reconstitution time
points. All available postchemotherapy immune data collected are shown
in Figure 1, with these data points representing various, rather than
consistent subjects over time. As shown in this figure, recovery of
total CD4+ T cells was very slow in surviving patients,
with median values remaining very low for at least 6 months after
completion of chemotherapy.
Postchemotherapy statistical analysis of T-cell regeneration is focused
on the 2 time points, wherein lymphocyte subset data are available from
all potentially evaluable subjects: 3 months after chemotherapy,
wherein lymphocyte subset data are available for 12 of 12 disease-free
patients, and at 12 months after chemotherapy, wherein lymphocyte
subset data are available for 8 of 8 disease-free patients. Several
points of evidence taken from these time points suggest that
thymic-dependent pathways, rather than peripheral expansion of mature T
cells contained within the PBPC grafts, were central to
CD4+ T-cell regeneration in this patient population. First,
although there was no correlation between age and CD4+
count at 3 months ( = 0.06, P = .97), there was an
inverse correlation between age and CD4 T-cell number at 12 months
( = 0.78, P = .04), similar to that reported
previously in chemotherapy recipients not treated with PBPC infusions
and which appears to relate to age-associated declines in thymic
regenerative capacity. Second, as shown in Figure 2, CD45 isoform
analysis revealed that, although CD45RO+CD4+
cells predominated during the months after infusion of PBSC products, CD4 counts did not normalize until normalization of CD45RA+
cells occurred.
Third, increased numbers of mature T cells within the PBPC graft did
not lead to enhanced CD4+ T-cell recovery. Indeed, at 3 months, the relationship between mature T cells infused and
CD4+ T cells was = 0.03, (P = .94) and at
12 months after therapy, there was no evidence for a positive
relationship between T-cell number infused and CD4+ number
( = 0.51, P = .18) (Figure
3). Further, because 10 of 19 patients
received CD34-selected grafts, direct comparison could be made between
patients who received T-cell-replete versus CD34 selected/T-cell-depleted products. The mean T-cell dose administered in the unmanipulated T-cell-replete grafts was
104 ± 40.5 × 106 cells/kg (mean CD4:CD8 ratio
1.8 ± 0.34) and the mean T-cell dose given to recipients of CD34
selected/T-cell-depleted grafts was
0.2 ± 0.04 × 106 cells/kg. Remarkably, as
shown in Figure 1, CD4+ T-cell reconstitution was
unaffected by whether T-cell-replete or T-cell-depleted PBPC grafts
were infused: median CD4+ count in T-cell-replete versus
CD34 selected/T-cell-depleted at 3 months 83 versus 84 cells/µL
(P = .75), 12 months 258 versus 445 cells/µL
(P = .65).
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| Fig 3.
Relationships between CD4+ T-cell
regeneration at 3 months and 12 months of age, CD34+ cell
numbers infused, and T-cell number infused at the time of
PBPC-supported chemotherapy.
For patients who received multiple cycles of PBPC-supported
chemotherapy, the mean CD34 and mean T-cell numbers received/cycle are
shown. Statistical significance is observed only for the relationship
between age and CD4+ T-cell count at 12 months
(P = .039) using the Spearman correlation.
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Because thymic-dependent pathways rely on an adequate supply of
prethymic progenitors, it is possible that enhanced T-cell regenerative
capacity in young patients reflects more vigorous CD34 mobilization.
However, we observed no correlation between the median
CD34+ cell number infused per cycle and age ( = 0.09,
P = .71), or the median CD34+ cell number infused
per cycle and CD4+ number 3 months ( = 0.21,
P = .29) or 12 months after therapy ( = 0.42,
P = .26) (Figure 3). Taken together, these results show that
provision of large numbers of CD34+ cells is not sufficient
to rapidly reconstitute CD4+ T-cell populations.
Furthermore, although the number of patients in each subset
is relatively small in this cohort, the results suggest that
the provision of significant numbers of autologous CD4+
T cells within the graft did not enhance CD4+
T-cell regeneration, whereas age-related changes within the host play a primary role in determining the rate of CD4+ T-cell
regeneration after chemotherapy.
In contrast to CD4 recovery, CD8 recovery was relatively brisk. As
shown in Figure 1, median CD8+ T-cell numbers were within
the normal range within 3 months of completion of chemotherapy in all
patients studied with some patients showing supranormal levels.
Although the lack of correlations in this small subset do not
conclusively rule out the possibility of biologically important
relationships that may exist, we observed no evidence for improved
CD8+ recovery in recipients of unmanipulated compared with
CD34 selected/T-cell-depleted grafts. Indeed, at 3 months after
infusion, recipients of CD34+ selected/T-cell-depleted
grafts had higher CD8+CD3+ counts (median 623 cell/µL [range 77-894]) compared with recipients of unmanipulated
grafts (median 188 [range 32-364], P = .055). There were no
significant differences in peripheral blood
CD8+CD3+ numbers in these 2 groups at 12 months
after therapy (median 707 [range 346-1685] in CD34 selected/T-cell
depleted versus 357 cells/µL [range 159-840] in unselected/T-cell
replete, P = .30). Similar distinctions in the pace of
CD4+ versus CD8+ T-cell recovery after
chemotherapy alone has been reported previously.11,12
To evaluate overall T-cell function in patients after chemotherapy,
functional proliferative assays were performed to analyze T-cell
responses to mitogens and recall antigens. As shown in Figure
4, there was a profound reduction in PHA
responses for a prolonged period after the completion of chemotherapy.
Similarly, all patients tested also showed a loss of responses to the
recall antigen tetanus toxoid during therapy, which persisted for at least 18 months after therapy (data not shown). Therefore, despite relatively rapid recovery of CD8+ T cells, these patients
remained significantly immunocompromised for a prolonged period after
PBPC infusions.
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| Fig 4.
Proliferative responses of peripheral blood mononuclear
cells to phytohemagglutinin during chemotherapy and after PBPC
infusions as described in "Materials and methods."
Net proliferative responses as measured in cpm are expressed as a ratio
compared with net proliferative response of PBMCs from a simultaneously
studied healthy donor. Pre-rx n = 13, Cy6 n = 15, 1 month n = 9,
3 months n = 9, 6 months n = 3, 9 months n = 4, 12 months
n = 4, and 24 months n = 2.
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B-cell and natural killer cell immune reconstitution
Median B-cell numbers returned to the low normal range by 3 months
after therapy and by 6 months after completion of therapy had
essentially normalized in all patients studied (Figure 1). Interestingly, several patients showed an initial overshoot in peripheral blood B-cell populations during the year after completion of
chemotherapy, similar to that observed with CD8+ T-cell
populations. With regard to serum immunoglobulin levels, intensive
multiagent chemotherapy induced statistically significant reductions in
all isotypes as shown in Figure 5, with the
most profound effects seen on serum IgM levels. Although many patients displayed a return to baseline values for serum IgG and IgA within 3 to
6 months after the completion of therapy, statistically significant reductions (P < .05 by paired sign rank test) in serum IgM
and serum IgA levels, compared with baseline values, were still
observed 9 months after the completion of therapy. These data
demonstrate that, although recovery of B-cell populations is brisk
after such intensive therapy, normalization of the antibody isotype
repertoire may not occur for a prolonged period.
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| Fig 5.
Serum immunoglobulin levels during sequential cycles of
chemotherapy, PBPC infusions, and after completion of therapy.
Shaded areas represent 95% confidence intervals for normal values. All
available immunoglobulin levels from the initial cohort of 19 patients
are shown. Evaluation of immunoglobulin levels was discontinued at the
time of tumor recurrence, therefore these data points represent
various, rather than consistent, subjects over time. Asterisks denote
median values that are significantly reduced from baseline using the
paired sign rank test (P < .05). The time points noted as 1 to 30 months represent time after completion of PBPC-supported
chemotherapy. Patients received 1 to 3 PBPC cycles as detailed in Table
1. Pre-rx n = 18, Cy3 n = 13, PBPC1 n = 15, PBPC2 n = 11, PBPC3
n = 11, 1 month n = 13, 3 months n = 8, 6 months n = 7, 9 months n = 8, 12 months n = 6, 15 to 18 months n = 4, and 24 months n = 2.
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As shown in Figure 1, there was a small, nonsignificant decline in
peripheral blood NK cell numbers when comparisons were made between
baseline cell counts and those obtained 1 month after the completion of
therapy (P = .07). However, it should be noted that there was
a great deal of interpatient variation in NK numbers, with some
patients displaying profound depletion and others retaining normal NK
cell number throughout the entire course of chemotherapy. These results
illustrate that, unlike CD4+ and B-cell populations, which
show profound depletion in all patients, the NK cell lineage is
relatively resistant to the effects of cytotoxic therapy in some
patients. Further, when NK cells were tested in the months immediately
following chemotherapy, there was evidence of lytic activity against
K562 targets in vitro (data not shown), providing evidence that this
lineage retained function as well.
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Discussion |
Age-associated thymic involution prevents rapid reconstitution of
CD4+ T-cell populations in a variety of clinical settings.
For instance, adults show a limited capacity for CD4+
regeneration post-BMT,13,14 whereas young patients show
rapid reconstitution of CD4+ T-cell numbers and T-cell
function.2,15 Similar age-related limitations in T-cell
regenerative capacity have been seen in the context of HIV infection,
in which thymic insufficiency has been invoked to play a role as
well.16,17 Moreover, it has been postulated that
age-related reductions in thymopoiesis play an important role in
immunologic defects associated with aging.18 However,
despite the importance of age-associated thymic involution for a
variety of clinical scenarios, the factors responsible for this process
remain incompletely understood.
Animal models have shown that profound reductions in the supply of
prethymic progenitors can result in thymic atrophy19 and
modest reductions in marrow-derived prethymic progenitors have been
shown in aging mice.20,21 This has led some investigators to postulate that age-associated thymic involution could result from
abnormalities in the supply of T-cell progenitors rather than due to
abnormalities within the thymus itself. In the setting of autologous
PBPC transplantation, one could postulate that cytotoxic effects of
chemotherapy and ablative transplant regimens on marrow-derived prethymic progenitors could limit the capacity for thymic-dependent T-cell regeneration in an age-dependent fashion. To investigate this
possibility, we evaluated T-cell regeneration in a population of
patients receiving large numbers of pluripotent hematopoietic progenitors in the context of dose-intensive chemotherapy. Although this was a relatively small cohort, we observed no relationship between
age and the number of CD34+ cells per kilogram administered
and no relationship between the dose of CD34+ cells infused
and subsequent T-cell regeneration. Furthermore, these patients
generally experienced rapid restoration of hematopoietic lineages as
well as B-cell, NK-cell, and even CD8+ T-cell populations,
suggesting that significant functional alterations in pluripotent
progenitor populations did not exist. However, a relationship between
age and CD4+ recovery was observed, similar to that
reported previously in patients receiving dose-intensive chemotherapy
without PBPC infusions. These results suggest that age-associated
constraints on CD4 recovery do not occur at the level of the
marrow. Similar results were recently reported using a model of BMT in
aged mice,22 wherein it was shown that the
provision of young bone marrow did not reverse age-associated thymic
involution. Taken together, the data suggest that
age-associated thymic involution is not primarily related to
an age-associated decline in the supply of marrow-derived prethymic progenitors, but rather is due to changes at the level of the thymus itself.
It is important to note, however, that in the allogeneic setting, PBPC
infusions provide more rapid CD4+ T-cell immune
reconstitution than BMT, suggesting some positive effect of the PBPC
graft in CD4+ T-cell regeneration. Indeed, Pavletic et
al5 reported significant reconstitution of CD4+
T-cell numbers as early as 28 days postallogeneic PBPC infusion, which
persisted for several months. Similarly, Ottinger et al6 reported more rapid and sustained recovery of CD4+ T cells,
and significantly higher CD4:CD8 ratios for up to 11 months after
transplant in recipients of allogeneic PBPC grafts compared with BMT
recipients. In these patients, normalization of responses to PHA was
observed as early as 3 months postallogeneic BMT. In contrast, in the
patients evaluated in this report, CD4+ T-cell numbers
remained very low in most patients until 9 to 12 months after therapy
and responses to PHA were diminished for at least 1 year post-PBPC
infusion. In a study of immune reconstitution after autologous BMT
versus autologous PBSC transplantation, Talmadge et al8
reported only a transient increase in CD4+ T-cell numbers
after PBSC transplantation compared with BMT at day 15, whereas no
differences were observed at later time points. Therefore, there appear
to be important differences between allogeneic and autologous PBPC
grafts with regard to their capacity to restore CD4+ T-cell populations.
Although such differences could relate to cytotoxic effects on
prethymic T-cell progenitors, the rapid restoration of other lineages
suggests that primitive cells are not functionally impaired. Alternatively, the data presented here suggest that T cells contained within the autologous graft may be significantly affected by cytotoxic therapy administered before the harvest. Indeed, in Figure 2 we show
that significant reductions in CD4+ T-cell numbers have
already occurred before the time of PBPC harvest. Not surprisingly
then, as shown in Table 1, even T-cell-replete grafts in this report
frequently contained fewer T-cell numbers than is commonly observed in
allogeneic PBPC grafts.9 Furthermore, at the time of PBPC
harvest, peripheral T cells showed evidence of widespread activation.
Activated T cells have been shown to undergo activation-induced cell
death after further stimulation through the T-cell receptor
(TCR).23,24 During immune reconstitution after
chemotherapy in adults, we have shown previously that stimulated CD4+ cells undergo apoptosis at a significantly higher rate
than CD4+ cells from normal controls. This elevated
susceptibility to apoptosis is correlated with increased expression of
the activation marker HLA-DR.12 In preliminary experiments
in the current study, a similarly elevated apoptotic rate was observed
in mitogen-stimulated CD4+ cells from selected patients
studied 3 to 12 months after therapy. In addition to a propensity for
activation-induced cell death, limitations in TCR repertoire diversity
may be predicted to exist in chemotherapy-depleted T-cell populations,
which could limit the capacity of mature T cells contained within the
graft to effectively contribute to immune reconstitution. Indeed,
Bomberger et al25 have shown significant limitations in TCR
repertoire diversity after autologous PBPC transplantation, both in the
presence and absence of T-cell depletion of the autologous PBPC graft.
The clinical implications of the profound, prolonged CD4 depletion
induced in this patient population are difficult to assess. Infectious
complications were common in this protocol with a greater than 20%
incidence of culture-proven bacteremia during chemotherapy cycles in
this patient population, which temporally coincided with episodes of
neutropenia. With regard to opportunistic and viral infections that may
be more directly related to T-cell depletion, 11 of the 19 patients had
complications develop that included P carinii
pneumonia,2 herpes zoster,1 Haemophilus
parainfluenzae pneumonia,2 mucocutaneous herpes
simplex,3 and mucocutaneous candidiasis2 and 1 case of clinically suspected, but unconfirmed Candida
endopthalmitis. In all cases, infectious complications were
successfully managed with supportive care and antimicrobial therapy.
More intriguing perhaps, is the potential role that such profound
T-cell depletion may have on the ability to eradicate residual tumor in
high-risk cancer patients. This report clearly demonstrates that tumor
recurrence was frequent and often preceded recovery of immune
populations. One possible means for improving outcome in patients with
high-risk tumors such as these is to augment cellular immunity against
specific tumor antigens.26-28 Indeed, immune-based
therapies are currently being explored for treatment of the pediatric
sarcomas that are the focus of this report.29 Animal models
have consistently shown, however, that such therapies are most likely
to be effective if undertaken in the setting of minimal residual
neoplastic disease.30,31 Clinical proof of concept of the
power of immune mediated responses to eradicate minimal residual
neoplastic disease has come from studies of graft versus leukemia in
the post-BMT setting. However, the data presented here clearly
illustrate that if immune-based therapies are to be applied in the
setting of minimal residual disease after autologous BMT, specific
measures must be taken to enhance immune reconstitution. On the basis
of our hypothesis that T-cell populations previously exposed to
chemotherapy are limited both quantitatively and functionally, one
possible strategy would be to harvest T-cell populations before cytotoxic therapy to be used as a source for CD4+ immune
reconstitution in the setting of minimal residual disease. Alternatively, future studies could involve the use of agents such as
IL-7, which have been shown in animal models to enhance T-cell
regeneration in vivo.32,33
In summary, we report persistent, profound CD4+ T-cell
depletion after high-dose chemotherapy, despite the administration of large numbers of CD34+ cells and in some patients large
numbers of T cells. These results illustrate that, in the autologous
setting, the age-associated decline in CD4+ T-cell
regenerative capacity is not reversed by the administration of a
sizable number of pluripotent hematopoietic progenitors. Furthermore,
because such autologous products are collected during administration of
intensive chemotherapy, we postulate that the replicative potential of
the mature T cells contained within the grafts is diminished, thus
limiting the capacity for such cells to contribute to peripheral
expansion pathways of CD4+ T-cell regeneration.
| |
Acknowledgments |
We acknowledge the clinical staff of the Pediatric Oncology Branch for
the excellent care rendered to patients treated on these protocols and
for their diligence in acquiring specimens for analysis. We would also
like to thank Dr Michael Bishop and Dr Seth Steinberg for their careful
reviews of the manuscript.
| |
Footnotes |
Submitted June 8, 1999; accepted March 7, 2000.
Reprints: Crystal L. Mackall, 10 Center Dr, MSC 1928, Bethesda,
MD 20892; e-mail: cm35c{at}nih.gov.
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.
| |
References |
1.
Mackall CL, Granger L, Sheard MA, Cepeda R, Gress RE.
T cell regeneration after bone marrow transplantation: differential CD45 isoform expression on thymic-derived versus thymic-independent progeny.
Blood.
1993;82:2585-2594[Abstract/Free Full Text].
2.
Weinberg K, Annett G, Kashyap A, Lenarsky C, Forman SJ, Parkman R.
The effect of thymic function on immunocompetence following bone marrow transplantation.
Biol Blood Marrow Transplant.
1995;1:18-23[Medline]
[Order article via Infotrieve].
3.
Storek J, Witherspoon RP, Storb R.
T cell reconstitution after bone marrow transplantation into adult patients does not resemble T cell development in early life.
Bone Marrow Transplant.
1995;16:413-425[Medline]
[Order article via Infotrieve].
4.
Mackall CL, Fleisher TA, Brown MR, et al.
Age, thymopoiesis and CD4+ T lymphocyte regeneration after intensive chemotherapy.
N Engl J Med.
1995;332:143-149[Abstract/Free Full Text].
5.
Pavletic ZS, Joshi SS, Pirruccello SJ, et al.
Lymphocyte reconstitution after allogeneic blood stem cell transplantation for hematologic malignancies.
Bone Marrow Transplant.
1998;21:33-41[Medline]
[Order article via Infotrieve].
6.
Ottinger HD, Beelen DW, Scheulen B, Schaefer UW, Grosse-Wilde H.
Improved immune reconstitution after allotransplantation of peripheral blood stem cells instead of bone marrow.
Blood.
1996;88:2775-2779[Abstract/Free Full Text].
7.
Roberts MM, To LB, Gillis D, et al.
Immune reconstitution following peripheral blood stem cell transplantation, autologous bone marrow transplantation and allogeneic bone marrow transplantation.
Bone Marrow Transplant.
1993;12:469-475[Medline]
[Order article via Infotrieve].
8.
Talmadge JE, Reed E, Ino K, et al.
Rapid immunologic reconstitution following transplantation with mobilized peripheral blood stem cells as compared to bone marrow.
Bone Marrow Transplant.
1997;19:161-172[Medline]
[Order article via Infotrieve].
9.
Korbling M, Huh YO, Durett A, et al.
Allogeneic blood stem cell transplantation: peripheralization and yield of donor-derived primitive hematopoietic progenitor cells (CD34+Thy-1dim) and lymphoid subsets, and possible predictors of engraftment and graft-versus-host disease.
Blood.
1995;87:2842-2848.
10.
Mackall CL, Fleisher TA, Brown MR, et al.
Lymphocyte depletion during treatment with intensive chemotherapy for cancer.
Blood.
1994;84:2221-2228[Abstract/Free Full Text].
11.
Mackall CL, Fleisher TA, Brown MR, et al.
Distinctions between CD8+ and CD4+ T cell regenerative pathways result in prolonged T cell subset imbalance after intensive chemotherapy.
Blood.
1997;89:3700-3707[Abstract/Free Full Text].
12.
Hakim FT, Cepeda R, Kaimei S, et al.
Constraints on CD4 recovery post chemotherapy in adults: thymic insufficiency and apoptotic decline of expanded peripheral CD4 cells.
Blood.
1997;90:3789-3798[Abstract/Free Full Text].
13.
Forman SJ, Nocker P, Gallagher M, et al.
Pattern of T cell reconstitution following allogeneic bone marrow transplantation for acute hematological malignancy.
Transplantation.
1982;34:967-998.
14.
Atkinson K, Hansen JA, Storb R, Goehle S, Goldstein G, Thomas ED.
T-cell subpopulations identified by monoclonal antibodies after human marrow transplantation. I. Helper-inducer and cytotoxic-suppressor subsets.
Blood.
1982;59:1292-1298[Abstract/Free Full Text].
15.
Buckley RH, Schiff SE, Schiff RI, et al.
Haploidentical bone marrow stem cell transplantation in human severe combined immunodeficiency.
Semin Hematol.
1993;30:92-101[Medline]
[Order article via Infotrieve].
16. Vigano A, Principi N, Bricalli D, et al. Immune reconstitution after
potent antiretroviral therapy in children with HIV infection
[abstract]. International Conference on the Discovery and Clinical
Development of Antiretroviral Therapies. 1998:75.
17.
Cohen Stuart JWT, Slieker WAT, Rijkers GT, et al.
Early recovery of CD4+ T lymphocytes in children on highly active antiretroviral therapy.
AIDS.
1998;12:2155-2159[Medline]
[Order article via Infotrieve].
18.
Hodes RH.
Aging and the immune system.
Immunol Rev.
1997;160:5-8[Medline]
[Order article via Infotrieve].
19.
Bosma GC, Custer RP, Bosma MJ.
A severe combined immunodeficiency mutation in the mouse.
Nature.
1983;301:527-530[Medline]
[Order article via Infotrieve].
20.
Fridkis-Hareli M, Abel L, Globerson A.
Patterns of dual lymphocyte development in co-cultures of foetal thymus and lymphohaemopoietic cells from young and old mice.
Immunology.
1992;77:185-188[Medline]
[Order article via Infotrieve].
21.
Tyan ML.
Age-related decrease in mouse T cell progenitors.
J Immunol.
1977;118:846-851[Abstract/Free Full Text].
22.
Mackall CL, Punt JA, Morgan P, Farr AG, Gress RE.
Thymic function in young/old chimeras: substantial thymic T cell regenerative capacity despite irreversible age-associated thymic involution.
Eur J Immunol.
1998;28:1886-1893[Medline]
[Order article via Infotrieve].
23.
Green DR, Bissonnette RP, Glynn JM, Shi Y.
Activation-induced apoptosis in lymphoid systems.
Semin Immunol.
1992;4:379-388[Medline]
[Order article via Infotrieve].
24.
Muro-Cacho CA, Pantaleo G, Fauci AS.
Intensity of apoptosis correlates with the general state of activation of the lymphoid tissue and not with stage of disease or viral burden.
J Immunol.
1995;154:5555[Abstract].
25.
Bomberger C, Singh-Jairam M, Rodey G, et al.
Lymphoid reconstitution after autologous PBSC transplantation with FACS-sorted CD34+ hematopoietic progenitors.
Blood.
1998;91:2588-2600[Abstract/Free Full Text].
26.
Velders MP, Schreiber H, Kast WM.
Active immunization against cancer cells: impediments and advances.
Semin Oncol.
1998;25:697-706[Medline]
[Order article via Infotrieve].
27.
Pardoll DM.
Cancer vaccines.
Nat Med.
1998;4:525-531[Medline]
[Order article via Infotrieve].
28.
Hellstrom I.
Tumor vaccines a reality at last?
J Immunother.
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Abstract
Introduction