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Blood, Vol. 93 No. 2 (January 15), 1999:
pp. 728-736
Monitoring Human Blood Dendritic Cell Numbers in Normal
Individuals and in Stem Cell Transplantation
By
D.B. Fearnley,
L.F. Whyte,
S.A. Carnoutsos,
A.H. Cook, and
D.N.J. Hart
From the Christchurch Hematology/Immunology Research Group and
Department of Hematology, Christchurch Hospital, Christchurch, New
Zealand; and the Mater Medical Research Institute, Brisbane,
Queensland, Australia.
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ABSTRACT |
Dendritic cells (DC) originate from a bone marrow (BM) precursor and
circulate via the blood to most body tissues where they fulfill a role
in antigen surveillance. Little is known about DC numbers in disease,
although the reported increase in tissue DC turnover due to
inflammatory stimuli suggests that blood DC numbers may be altered in
some clinical situations. The lack of a defined method for counting DC
has limited patient studies. We therefore developed a method suitable
for routine monitoring of blood DC numbers, using the CMRF44 monoclonal
antibody (MoAb) and flow cytometry to identify DC. A normal range was
determined from samples drawn from 103 healthy adults. The mean
percentage of DC present in blood mononuclear cells (MNC) was 0.42%,
and the mean absolute DC count was 10 × 106 DC/L blood.
The normal ranges for DC (mean ± 1.96 standard deviation [SD]) were
0.15% to 0.70% MNC or 3 to 17 × 106 DC/L
blood. This method has applications for monitoring attempts to mobilize
DC into the blood to facilitate their collection for immunotherapeutic
purposes and for counting blood DC in other patients. In preliminary
studies, we have found a statistically significant decrease in the
blood DC counts in individuals at the time of blood stem cell harvest
and in patients with acute illnesses, including allogeneic bone marrow
transplant (BMT) recipients with acute graft-versus-host disease
(aGVHD).
© 1999 by The American Society of Hematology.
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INTRODUCTION |
DENDRITIC CELLS (DC) originate from bone
marrow (BM) precursors and circulate via the blood to most tissues of
the body where they have a sentinel role in immune surveillance. These peripheral DC acquire antigens and migrate via the lymph to regional lymph nodes where they initiate T-lymphocyte responses.1,2 DC populations may be heterogeneous and several potential
differentiation pathways have been described.2 Myeloid DC
in man are best identified by a combination of morphologic, phenotypic,
and functional features: there is at present no evidence for a human
lymphoid-derived blood DC. No surface marker specific for blood
myeloid-derived DC (hereafter DC), which are found among a
heterogeneous population of lineage marker negative (lin-)
HLA-DR positive cells, generally 1% to 2% of blood mononuclear cells
(MNC), has been described to date. As yet, no DC deficiency state has
been described in man, and there is little information on blood DC
numbers in disease. However, the turnover of DC in the tissues is
substantially increased by various inflammatory stimuli,3-6
and blood DC numbers might be raised to support this. Alternatively,
other BM-derived cells, for example monocyte-derived DC
(Mo-DC)7,8 might account for the increased turnover of DC
at sites of inflammation.
Previous studies have estimated blood DC numbers by extrapolating from
the yield of DC obtained (after multistep purification procedures) from
a given volume of blood. These estimates of DC number are dependent on
the isolation methods used and the criteria used to define DC, and thus
vary considerably. Moreover the enumeration of DC after the use of
complex isolation techniques is not applicable to the routine
monitoring of DC numbers. Early work estimated blood DC numbers as
being in the order of 0.1% to 0.5% of MNC,9 later quoted
as 0.3% of MNC.10 Subsequent estimates of blood DC number
range from approximately 0.1% of MNC11 to 0.9% to 1.0%
of MNC,12,13 although higher estimates (4% of
MNC14) have been published. DC numbers appear to be
decreased in the blood of acquired immunodeficiency syndrome (AIDS)
patients13 and in patients with advanced stages of breast
cancer.15 The ability to measure DC numbers readily in
patients would encourage further studies of the role DC play in disease
and facilitate attempts to mobilize DC into the blood to be collected
for immunotherapeutic purposes.
There are precedents for counting rare cell populations by flow
cytometry, notably the routine monitoring of blood CD34+
cell numbers.16,17 Attempts to measure DC number are
complicated by the lack of a suitable specific marker for circulating
blood DC.18,19 However, blood DC do express some relatively
restricted activation antigens, notably the CD8320,21 and
CMRF4422,23 antigens, after a brief period of in vitro
culture. The CMRF44+/CD14 and CD19 cells
generated in this manner have the morphology, phenotype, and functional
attributes of DC.23 The relatively rapid upregulation of
the CMRF44 antigen23 indicated that it might be possible to
use the CMRF44 monoclonal antibody (MoAb) to measure and monitor blood
DC numbers. We now report a flow cytometric method, which uses the
CMRF44 MoAb to count the DC, after blood MNC have been activated by a
brief period of in vitro culture. The method has been used to determine
a normal range for blood DC counts in a healthy adult population. As a
preliminary study of the method's use in a clinical setting, we
measured blood DC counts in patients undergoing blood stem cell
mobilization before transplantation and monitored DC counts in patients
undergoing autologous and allogeneic blood stem cell or bone marrow
transplantation (BMT). The resulting data supports the hypothesis that
blood DC numbers may be altered in certain disease states.
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MATERIALS AND METHODS |
Subject selection.
A reference population was established from healthy volunteers aged 19 to 86 years, who had had no recent illness at the time of specimen
collection. All samples were collected between 8 am and 2 pm. Patients
undergoing autologous or allogeneic stem cell transplantation in this
institution were invited to contribute to this study. Blood samples
were taken after engraftment (neutrophils >1.0 × 109/L). Other patients with a single disorder were also
tested (Table 1). BM was obtained by
aspiration from the posterior iliac crest of volunteer normal donors.
Cord blood was collected by venipuncture of the umbilical vein of
normal full-term deliveries. All specimens were obtained with the
approval of the local Ethical Committee after informed consent had been
given by the donor (or mother in the case of cord blood collections).
The stem cell mobilization protocol involved a single dose of
cyclophosphamide (2 g/m2) and granulocyte
colony-stimulating factor (G-CSF; Roche, Auckland, New
Zealand), 5 to 10 µg/kg/day from day 0 to
the end of harvesting, or G-CSF (10 µg/kg/day) alone. Harvests were
usually initiated on the day that the CD34+ count rose over
20 × 106/L.
Preparation of sample.
A total of 5 mL of blood was collected into EDTA and processed within 4 hours of collection as follows: a routine blood count was performed
using a Coulter STKS (Coulter Electronics, Hialeah, FL). The absolute
number of MNC/L blood was determined from this measurement. The
remaining blood was diluted 1:1 with sterile phosphate-buffered saline
(PBS) and underlayed with Ficoll-Paque (Pharmacia, Uppsala, Sweden)
before centrifugation at 520g for 15 minutes at room
temperature. The MNC were recovered from the low-density interface and
washed three times before culturing for 24 hours in RPMI-1640/10%
fetal calf serum (FCS) (Life Technologies, Auckland, NZ) at 37°C
and 5% CO2 in air in Falcon 2054 round bottom polystyrene
tubes (Becton Dickinson, Sydney, Australia). Typically, a 5 mL blood
sample yielded approximately 5 × 106 MNC, which were
cultured at a concentration of 1 × 107 cells/mL
(normally 0.25 to 1 mL media). In some experiments, cells from
individual donors were cultured for 4, 24, and 48 hours. Granulocyte-macrophage (GM)-CSF (Sandoz, Basel, Switzerland), interleukin-3 (IL-3) (Sandoz), and tumor necrosis factor (TNF) (Hoffman-La Roche, Basel, Switzerland) were used at the indicated concentration to document the effect of adding cytokines to this culture media. RPMI/10%FCS was chosen as the culture media on the
basis of a previous comparison of the ability of different media to
support DC in vitro24 and to provide standard culture conditions in individuals in whom drugs or other serum factors may have
affected DC activation. After culture, cells were washed and their
viability checked using trypan blue exclusion. A repeat blood sample
was requested in the rare instances where the viability was <95%.
Cells were then resuspended at 107/mL and labelled as
described below.
MoAbs.
The MoAb, CMRF44 (IgM)22, was produced in this laboratory,
as was the IgM negative control MoAb, CMRF50, which recognizes a
keratinocyte antigen and does not react with hematopoeitic cells. Phycoerythrin (PE)-conjugated CD14 (leuM3, IgG2a), CD19 (leuM12, IgG1),
anti-HLA-DR (L243, IgG2a), and IgG2a negative control MoAb were
purchased from Becton Dickinson. CD83 (HB15a, IgG2b) was purchased from
Immunotech (Marseille, France). Fluorescein isothyocyanate-conjugated sheep antimouse immunoglobulin (FITC-SAM) was purchased from Silenus (Hawthorne, Victoria, Australia). Propidium iodide (PI) was purchased from Sigma (St Louis, MO). CMRF31 (CD14, IgG2a, this laboratory), FMC63
(CD19, IgG1, a gift of Prof H. Zola, Adelaide, Australia), HuNK2 (CD16,
IgG2a, a gift from Prof I. McKenzie, Melbourne, Australia), OKT3 (CD3,
IgG2a, from a hybridoma obtained from the American Type Tissue
Collection (Rockville, MD) were used to create a MoAb cocktail against
the B and T lymphocyte, monocyte and natural killer (NK) leukocyte
lineages. Cells that did not label with this MoAb cocktail were
designated lin.
Labelling procedure.
Cells were labelled using standard techniques. Briefly, 50 µL of
cells (107/mL) were placed into five tubes. Two tubes were
labelled with the negative control MoAb and three with the CMRF44 MoAb
for 20 minutes at 4°C, before being washed twice in PBS/1%bovine
serum albumin (BSA) and then labelled with FITC-SAM for another 20 minutes. After further washing and blocking using 10% mouse serum,
PE-conjugated CD14 and CD19 MoAb were added to each tube and incubated
for 20 minutes at 4°C. A final wash was followed by the addition of
5 µL of 10 µmol/L PI to each tube.
Flow cytometry.
Analysis was performed using a fluorescence-activated cell sorting
(FACS) vantage flow cytometer (Becton Dickinson, San Jose, CA) equipped
with a 488 nm argon ion laser with fluorescence emission detected at
525, 575, and 675 nm. A total of 50,000 events was acquired from each
tube and analyzed using Cellquest software (Becton
Dickinson). Live cells were gated on the basis of forward and side light scatter characteristics and PI exclusion. A region, which encompassed CMRF44+ CD14
CD19 events, was copied onto all five dot plots. On the
negative control dot plots, this region typically contained <0.1% of
total events. The percentage of CMRF44+ CD14
CD19 events was noted and the mean of the two negative
control values was subtracted to give triplicate measurements of DC
number as a percentage of total MNC. The mean of these triplicate
values was taken as the final DC percentage of MNC and used to
calculate the absolute DC number.
Statistics.
Statistical analysis (t tests for independent groups and paired
data as appropriate) was performed using the Instat statistics package
(GraphPad Software, San Diego, CA).
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RESULTS |
Analysis of blood DC numbers by flow cytometry.
The kinetics of CMRF44 expression on cultured lin, HLA
class II+ cells (enriched by magnetic depletion of
lin+ cells) was checked to determine the optimal period of
in vitro culture for this test. The CMRF44 antigen is one of the first activation antigens expressed by DC,23 but culture for less than 6 hours was insufficient to allow accurate detection of DC (Fig 1A). Culture times of 12 hours clearly
distinguished CMRF44+ cells, but are not convenient for a
routine laboratory. The number of CMRF44+ DC was near
maximal after 16 to 48 hours. Although the mean fluorescence intensity
of the positive cells was slightly higher at 48 rather than 24 hours,
the viability of the cultured cells was better at 24 hours and
therefore 24 hours in vitro culture was selected as a suitable culture
period for routine DC counting.
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| Fig 1.
Time course of CMRF44 antigen expression (normalized,
100% = maximum number of cells labelling with CMRF44) on: (A)
lin (DC enriched) sorted cell populations cultured for
the period indicated (n = 6). (B) Blood MNC, cultured as described
for routine DC counting (solid line, normal individuals (n = 3),
dashed line, patients (n = 3).
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Flow cytometric analysis of the MoAb-labelled cells enabled the
percentage of DC present in cultured MNC populations to be measured
(see Materials and Methods and Fig 2).
Events that were CMRF44+ CD14
CD19 (R3 in Fig 2C) were, after subtraction of the
control (background staining cells), counted as DC and expressed as a
percentage of the blood MNC. Absolute DC counts were then calculated
from the number of MNC/L blood (as determined by the automated cell
counter), multiplied by the percentage of DC. Absolute DC counts varied independently of the number of MNC and did not correlate with either
the absolute monocyte or lymphocyte counts (r2 = 0.05 and
0.06, respectively).
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| Fig 2.
FACS analysis of labelled cells. (A) MNC were gated (R1)
and dead cells excluded by PI labelling (R2 not shown). A total of
50,000 events was aquired from each tube. (B) The number of
CD14 and CD19 events occuring with the
negative control antibody was measured (R1 and R3). Two negative
control tubes were analyzed, and the mean of these two values (eg,
0.04% in the example shown) was used. (C) Events in R1 and R3
(CMRF44+ CD14 and CD19
cells) were counted in three separate tubes. The mean negative control
value (B) was subtracted from each value. The mean of these three
replicates was used as the DC percentage of MNC. The absolute DC count
was calculated as described in the Results.
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The precision of the method was tested by performing multiple
simultaneous tests on individual donors. The variance of the three
replicates of the single tests performed on 97 individuals was 0.004%.
When three simultaneous tests of three replicates were performed on
three normal donors, the variance of the replicates was 0.004% with an
additional component of variance due to the tests of 0.0036%. This
suggested that other test factors, as well as the labelling
and/or analysis of the replicate tubes, accounted for most of
the observed variance, but that other factors also contributed to the
variance. Therefore, the effect of varying the MNC concentration during
the culture and the effect of titrating conditioned media into the cell
culture was studied. Variation between the conditions tested in this
manner appeared no greater than that seen between replicates performed
in identical conditions (data not shown), suggesting that labelling
and/or analysis accounted for most of the observed variation.
However, when normal individuals were tested at intervals of >1 week,
considerable variations in relative and absolute DC numbers were seen (Table 2). This may
reflect fluctuation within a relatively broad range, as is seen with
other leukocytes, or could be due to cyclical variation in blood DC
numbers. It was unlikely to be due to circadian variation, as all
samples were collected at a similar time of the day.
Analysis of the reference population and determination of a normal
range.
The reference population comprised 103 normal individuals, drawn mainly
from a population of hospital and laboratory workers, but also the
general public. The age range of this population was 19 to 86 years and
the male:female ratio was 50:53. The mean number of DC as a percentage
of MNC was 0.42% (95% confidence interval [CI] = 0.39 to 0.45) and
the observed range was 0.12 to 0.94% MNC. The mean absolute DC number
was 9.8 × 106/L blood (95% CI = 9.1 to 10.4), and
the observed range 3 to 26 × 106 DC/L
(Fig 3A). These results showed an
approximately normal distribution (Fig 3B).
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| Fig 3.
(A) Scatter plot of the distribution of DC numbers in 103 normal individuals, expressed as percentage of blood MNC and as
absolute counts. (B) DC numbers appear to follow a normal distribution.
The mean DC proportion (0.42% of MNC) and the calculated normal range
(0.15% to 0.70% of MNC) are shown (solid and dashed lines,
respectively). Absolute DC counts follow a similar distribution (not
shown).
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The mean percentage and absolute DC number did not differ significantly
when females and males were analyzed separately, although the range was
slightly greater in females aged under 50, raising the possibility that
hormonal factors may influence DC number. DC numbers did not vary
significantly when the reference population was segregated on the basis
of age and appeared to be maintained even in the oldest (13 patients > 60 years) subjects in this study.
A normal range, based on a mean of 0.42 ± 1.96 SD was derived as
0.15 to 0.70% of MNC and 3 to 17 × 106 DC/L blood.
Validation of the method.
It was theoretically possible that in clinical situations, factors such
as inflammatory mediators might affect CMRF44 antigen expression, and
thus the assay. Indeed, our initial experience was that DC numbers in
patients were often low. Therefore, additional experiments, including
the use of alternative methods for estimating DC number, were performed
to validate the method. Time course experiments were performed on
normal subjects and patients with inflammatory conditions to ensure
that the kinetics of CMRF44 expression was similar when unseparated MNC
were cultured for DC counting. A period of 20 to 24 hours was confirmed
in these individuals as suitable for DC counting, based on CMRF44
antigen expression, and the kinetics of its expression appeared similar in both the normal and patient groups (Fig 1B).
Supplementing the culture media with cytokines known to promote DC
survival (GM-CSF, TNF, or IL-3) did not significantly increase DC
numbers (Table 3), and the effects were
variable between individuals. In some cases these cytokines
(particularly the TNF) increased CRMF-44 and decreased CD14 or CD19
expression on non-DC populations, making clear discrimination of
CMRF44+/CD14 and 19 cells difficult.
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Table 3.
DC Counts Obtained in Five Individuals After Culture in
Media With Additional Cytokine Compared With Those Obtained Without
Supplementation
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Alternative methods of estimating blood DC number were used in parallel
with the CMRF44-based method to obtain some comparative data. The
number of lin/HLA-DR+ cells present before
cell culture was tested in 11 normal individuals and 13 patients with a
low mean DC count. A lower number of
lin/HLA-DR+ cells (0.60% v 0.85%)
paralleled the lower CMRF44+ DC number (0.21% v
0.49%) in these patients compared with the normals
(Fig 4A). Nonetheless, the relatively
constant relationship between the lin
HLA-DR+ cells and CMRF44+ DC seen in the normal
individuals was lost for the patients (Fig 4B). This probably reflected
the variable cellular composition of the lin
HLA-DR+ population.
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| Fig 4.
Comparision of DC numbers identified as either
lin HLA-DR+ cells (before culture) or as
CMRF44+ CD14 CD19 cells
(after overnight culture). (A) The mean numbers of
linHLA-DR+ cells and CMRF44+
DC were both reduced in patients (n = 13) compared with normal
individuals (n = 11). (B) The relatively constant relationship
between the numbers of CMRF44+ DC and
linHLA-DR+ cells seen in normal
individuals (where CMRF44+ DC typically account for 40%
to 80% of lin HLA-DR+ cells) is not seen
in the patient group (CMRF44+ DC account for 5% to 95%
of the lin HLA-DR+ cells). (C) The use of
CMRF44 or CD83 to identify DC was compared in both normals (n = 5)
and patients (n = 16). A strong correlation (r2
= 0.89) was found between the DC numbers measured using CMRF44 or
CD83.
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In 21 experiments (5 normal individuals and 16 patients), additional
labelling with an alternative DC activation marker, CD83, was
performed. The DC number was usually slightly (mean 15%) lower when
measured using this antibody, possibly as a result of the different
kinetics of CD83 antigen upregulation.23 There was a strong
correlation (r2 = 0.89) between CMRF44+ and
CD83+ DC numbers (Fig 4C) in both normal individuals and
patients. These results indicate that the low DC number seen in some
patients was not simply a consequence of altered CRMF44 expression
during illness.
Monitoring DC numbers during stem cell mobilization and harvesting.
Blood stem cell mobilizations were studied in 20 individuals; in 17 patients cyclophosphamide/G-CSF was used and in one patient and two
normal donors, G-CSF alone was used to mobilize stem cells into the
blood (Fig 5). DC numbers were assayed on
day 1 of the harvest in all patients. Where possible pre-
and/or postmobilization counts were done, although due to
logistical reasons, not all patients could be tested at all time
points. The mean percentage DC and absolute DC counts were
significantly lower than normal on day 1 of harvesting (DC percentage
0.14% v 0.42% of MNC, P < .01 and absolute DC
counts 2.9 v 9.8 x 106/L, P < .01). Using
all possible data points, paired analysis of DC numbers was also
performed and again the difference between the pre- and during harvest
DC number was significant (n = 11, 0.42 v 0.13% of MNC,
two-tailed P < .01: paired Student's t test) and
(8.0 v 2.6 × 106/L, P < .01) (Fig 5B and C). Two normal donors were harvested for allografts
after mobilization with G-CSF alone and a similar pattern was seen,
with a decrease in the DC blood count at the time of harvest. An
increase in DC count (some within the normal range) was noted 1 to 2 weeks after harvesting in all of the donors tested (Fig 5).
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| Fig 5.
Blood DC numbers at the time of stem cell mobilization in
20 subjects. (A) Mean DC percent of blood MNC before, during, and after
stem cell harvest. (B) Paired (pre- and during or during and post)
percent DC of blood MNC in individual patients. (C) Paired (pre- and
during or during and post) absolute DC counts. The differences in DC
counts before and during the harvest are statistically significant (see
text).
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For comparison, DC numbers in BM aspirates and cord blood were also
measured using this method. Normal BM aspirate DC numbers appeared
similar to normal blood DC numbers (mean marrow DC number = 0.41%, n = 3). The DC numbers in cord blood were lower than those seen in adult
blood (mean = 0.23%, n = 8), as has been reported previously.25
Analysis of blood DC counts after stem cell transplantation.
The range of DC counts after stem cell transplantation did not follow a
normal distribution (Fig 6). A total of 35 patients were studied, with 11 being tested on more than one occasion. Subdividing patients on the basis of graft type showed a similar mean
percentage of DC in both the allogeneic and autologous recipients. In
the autografted group, no difference was seen between DC numbers in
patients who received blood stem cell autografts (with low DC
percentages as above), CD34+ selected blood stem cell
autografts (with very low absolute DC numbers), or the patients who had
received a BM graft.
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| Fig 6.
Post-BMT DC counts from 35 patients (filled symbols) were
subdivided on the basis of the three criteria indicated. The mean DC
number for each patient group is shown (solid line). The mean (dashed
line) and the limits of the normal range (dotted lines) are shown for
comparison. Mean DC numbers were similar after autologous and
allogeneic grafts. The mean DC number for the patients with aGVHD was
significantly lower than the other postallogeneic stem cell transplant
patients (see text).
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In contrast, the percentage and absolute DC number were low in patients
with acute graft-versus-host disease (aGVHD) compared with other allo
BMT patients (mean DC number 0.17% v 0.54%, P = .01),
the only exception being in one patient with grade 1 skin aGVHD. A low
DC number (0.06% of MNC) was also seen in one pediatric patient with
aGVHD, but this result was not included in the above analysis due to
the lack of a formal normal range for this age group. The low DC number
is likely to be related to aGVHD rather than other associated factors,
as mean DC counts in the early (28 to 60 days) or later (>60 days)
posttransplant period did not differ significantly, and likewise mean
DC counts were similar after grouping patients on the basis of
cyclosporin A treatment (not shown). As steroids, usually prednisone or
dexamethasone, were used to treat aGVHD, these agents may have
contributed to the low DC counts in these patients, however chronic
GVHD (cGVHD) patients who were also receiving steroids showed no
obvious effect on DC numbers. In addition, the effect of high dose
steroids on DC number was monitored in two (nontransplant) patients
with immune thrombocytopenia, and no change in DC number was seen (not
shown).
Some patients with no intercurrent illness (including those with
recently resolved aGVHD) had high DC counts in the first 3 months
posttransplant, although the low MNC numbers in some patients meant
this did not always translate into a high absolute DC count.
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DISCUSSION |
Ideally, blood myeloid DC number would be determined with the use of a
marker specific for circulating DC. Because no such marker is currently
available, culture of MNC is required to allow the use of DC activation
antigens for positive identification of DC. The decision to use CMRF44
as the reference antibody for this study was based on the rapidity of
the upregulation of the CMRF44 antigen and its relative density of
expression on DC. These two factors are important in that they allow a
short period of in vitro culture with subsequent clear identification
of the DC by flow cytometry. CD83 can be used to enumerate blood
DC,11 but in our experience, CD83 antigen expression lags
behind CMRF44 antigen expression on DC and is not as strongly
expressed.23 Both of these factors may contribute to the
slightly lower DC numbers we observed when we used CD83 in place of
CMRF44. DC enumeration using CD83 after a multistep purification
suggested that DC numbers were in the range 0.06% to 0.15% of blood
MNC,11 but potential DC loss during purification, together
with the factors mentioned above, may account for these values, which
are lower than our range of 0.15% to 0.70% of blood MNC. In our
experience, DC yields after purification are often only approximately
50% of that expected based on blood DC numbers found in this
study.24 An alternative method based on a
lin HLA-DR+ DC phenotype has also been used
to directly enumerate blood DC,12 and the numbers of
lin HLA-DR+ we observed in normal
individuals (0.85%) correlated closely with the lin
HLA-DR+ cell number reported (0.90%). This approach has
the benefit of avoiding the requirement for cell culture, but is less
precise in its definition of a DC. The
linHLA-DR+ population is not homogeneous in
its expression of a number of surface markers and has been reported to
give rise to a number of different cell populations.26-28
Furthermore, expression of HLA-DR on other activated cells during
illness may also complicate the issue. Our results indicate that the
non-DC, lin HLA-DR+ populations may vary
considerably in number within patients, creating potential for error
when this method is used to monitor blood DC.
The need to culture cells introduces a number of potential variables,
which may influence the result and certain assumptions are inherent to
this method. First, it must be assumed that DC are not over or
under-represented in the cell losses, which occur during density
gradient separation and culture of the MNC. This method involves
minimal cell handling, and we excluded samples with low viability from
analysis to limit potential variance from this source. A second
assumption made in comparing DC numbers between populations is that the
DC number is not affected by the relative composition of the cultured
MNC. The relatively minor effect that cytokine supplementation of the
culture media had on DC numbers suggests that MNC cultured in RPMI/10%
FCS can provide a suitable millieu for DC maturation and survival over
the culture period used in this method. Validation of this method is
difficult in the absence of a reference standard, but these results
appear internally consistent and our mean of 0.42% MNC is similar to 0.47% (n = 8) reported by Olweus et al.29 The potential
confounding factors in our method have been identified and eliminated
as far as possible. The observed range in DC numbers in normals and
patients is relatively small (<1% of blood MNC), therefore we
derived a normal range from over 100 individuals to ensure random
variation did not unduly influence our results. This will provide a
useful reference standard for further refinement of DC counting
methods. Such advances are likely to include direct cell counting on
the analysis tube using comparative standards and direct labelling of
the DC with improved CMRF44 conjugates.
The clinical relevance of altered blood DC numbers is not yet clear
and, given the difficulty in determining the cause of the alterations
in blood DC number we documented, it is important not to overinterpret
these results. Variation in DC blood transit time may be important in
regulating the supply of DC to the tissues, yet may not alter the
number of DC present in the blood at a single point in time. The
information on blood DC kinetics needed to analyze these results is not
yet available, although studies on blood monocyte kinetics suggest that
normal monocyte blood transit times (half-life in circulation of 8 hours) are reduced during acute illness.30,31 The short
circulation time of monocytes and lymphocytes32 and the
rather low number of DC (approximately 5 × 107) in
the blood at any one time (balanced against the relatively large
estimated daily turnover of tissue DC5,6) all suggest that
the DC blood transit time is likely to be short.
Several factors may contribute to the low blood DC counts we observed
during acute inflammation. DC production and release into the blood may
be exceeded by an increased rate of egress of DC from the blood, either
at the site of inflammation or generally due to DC activation in the
bloodstream. Production of other myeloid cells during acute
inflammation may occur at the expense of DC production, either as a
direct consequence of a bipotential precursor cell becoming committed
to non-DC pathways33,34 or by inhibition of DC development
by other cell types or cytokines.35 These factors may
explain the fact that we found low blood DC counts in situations
similar to those in which other investigators (working mainly with
animal models) have found increased tissue DC turnover. Alternatively,
tissue DC turnover may be supported from an alternative source, perhaps
akin to the in vitro generation of Mo-DC.7,8 We did not
find DC counts to drop with age or steroids despite suggestions from
animal studies that these might affect DC numbers in the
tissues.36,37
This method will be useful for monitoring attempts to mobilize DC into
the blood. The numbers of other leukocyte types are generally reported
to remain constant or increase in G-CSF-mobilized blood.38,39 Our data suggests that DC numbers are reduced
at the time that CD34+ cells are harvested after
cyclophosphamide/G-CSF or G-CSF stimulation and augments previous data
on the composition of blood stem cell grafts (reviewed in To et
al40). This observation may explain, in part, the
previously reported finding that after G-CSF mobilization, blood MNC
are less allostimulatory on a cell for cell basis.41 However, the harvested product still contains a substantial number of
DC. There was no reason, apart from the precedent of the dramatic but
unexpected effect of G-CSF on stem cells, to suspect that DC numbers
would be altered by this commonly used stem cell mobilization protocol.
Interestingly, there appears to be little difference in DC numbers when
GM-CSF is used instead of G-CSF in the mobilization protocol (R. Brown,
personal communication, November 1992), despite the
well-documented effect of GM-CSF on in vitro DC
development.42,43 The kinetics of blood DC production in
this situation may be slower than other leukocytes, however, it is also
possible that G-CSF either directly or indirectly influences DC
production or transit times. These influences may also explain the
apparent increase in DC counts subsequent to harvesting.
A number of cytokines (principally combinations involving GM-CSF, IL-4,
and TNF, but also other factors8,43-46) are known to
promote the in vitro development of DC from several blood precursor cell populations. As yet, stimuli, which may specifically mobilize DC
into the blood, have not been described. The ability to harvest significant numbers of DC directly from the blood may offer a useful
alternative to the in vitro generation of these cells for clinical use.
In mice, flt3 ligand has been reported to increase DC numbers in
several organs, including the blood.47 Flt3 ligand effects
on human DC are now being examined and early results suggest DC are
increased in the blood.48 As experience with
CD34+ stem cell mobilization has shown, testing regimens
specifically designed to increase blood DC number will require a
practical and reproducible method of monitoring the cells in question,
and the methods used are likely to evolve over time. The method
reported here is practical and the small volume of blood required makes daily testing feasible. These results may be a useful basis for comparison as interest in DC mobilization for cancer immunotherapy increases.
Previous studies have shown that DC repopulation occurs rapidly after
BMT.49 Normal DC numbers were found in patients grafted with CD34+ selected cells, providing evidence that
CD34+ selected cells are capable of repopulating the DC
lineage. The fact that relatively high DC numbers were observed in
patients who were otherwise well after stem cell mobilization or
transplantation could be seen as a rebound phenomenon and may relate to
the previously reported finding that DC precursors (colony-forming
unit-dendritic Langerhans cell [CFU-DL]) are more numerous in blood
as counts recover after chemotherapy.50 In some of the
allogeneic BMT cases, skin biopsy material was available and our
preliminary data suggest that blood and skin DC numbers are both low in
aGVHD (Mckenzie, manuscript in preparation). Taking the
low DC numbers seen in aGVHD at face value, it may be that the initial
low DC number is due to increased turnover and is later depressed as a
result of decreased production of DC, possibly as part of a normal
immunoregulatory process. As we have also found low blood DC numbers in
four of five non-BMT patients with acute inflammation due to infections
or trauma, it is possible that this may be a general response to
inflammation and not an aGVHD specific effect.
The availability of this routine method will allow blood DC counts to
be correlated with clinical outcomes, possibly providing data that can
address the question of whether or not variations in DC number relate
to disease pathogenesis. The method and normal range we report here
represent an initial step towards achieving this goal. The initial
results obtained using this method suggest that monitoring DC counts
may be clinically useful, as well as providing data, which improves our
understanding of basic DC biology. The test has immediate application
for monitoring the production of DC for experimental immunotherapy
protocols.
| |
ACKNOWLEDGMENT |
We wish to thank the volunteers and patients who provided blood samples
and acknowledge the contribution of clinical colleagues and Alison
Inder in collecting patient specimens. Valuable statistical advice was
provided by Dr Elisabeth Wells. We thank Jan Merry-Martin for typing
the manuscript.
| |
FOOTNOTES |
Submitted March 23, 1998;
accepted September 15, 1998.
Supported by the New Zealand Health Research Council, Canterbury
Medical Research Foundation, Cancer Society of New Zealand, Lottery
Health Research Council, McClelland Trust, Leukemia and Blood
Foundation of New Zealand, Loch Maben Trust, University of Otago, and
others.
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.
Address reprint requests to Prof D.N.J. Hart, PhD, MD,
Director, Mater Medical Research Institute, Raymond Terrace, Brisbane,
Queensland, 4101 Australia; e-mail: dhart{at}mater.org.au.
| |
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Top
Abstract
Introduction