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Blood, 1 October 2000, Vol. 96, No. 7, pp. 2511-2519
IMMUNOBIOLOGY
Concept of lymphoid versus myeloid dendritic cell lineages
revisited: both CD8  and CD8+
dendritic cells are generated from CD4low
lymphoid-committed precursors
Pilar Martín,
Gloria Martínez del
Hoyo,
Fabienne Anjuère,
Sara Ruiz Ruiz,
Cristina Fernández Arias,
Alvaro Rodríguez Marín, and
Carlos Ardavín
From the Department of Cell Biology, Faculty of
Biology, Complutense University, Madrid, Spain.
 |
Abstract |
Two dendritic cell (DC) subsets have been identified in the murine
system on the basis of their differential CD8 expression. CD8+ DCs and CD8 DCs are considered as
lymphoid- and myeloid-derived, respectively, because
CD8+ but not CD8 splenic DCs were
generated from lymphoid CD4low precursors, devoid of
myeloid reconstitution potential. Although CD8 DCs
were first described as negative for CD4, our results demonstrate that
approximately 70% of them are CD4+. Besides
CD4 CD8 and CD4+
CD8 DCs displayed a similar phenotype and T-cell
stimulatory potential in mixed lymphocyte reaction (MLR),
although among CD8 DCs, the CD4+ subset
appears to have a higher endocytic capacity. Finally, experiments of DC
reconstitution after irradiation in which, in contrast to previous
studies, donor-type DCs were analyzed without depleting
CD4+ cells, revealed that both CD8+ DCs and
CD8 DCs were generated after transfer of
CD4low precursors. These data suggest that both
CD8+ and CD8 DCs derive from a common
precursor and, hence, do not support the concept of the
CD8+ lymphoid-derived and CD8
myeloid-derived DC lineages. However, because this hypothesis has to be
confirmed at the clonal level, it remains possible that CD8 DCs arise from a myeloid precursor within the
CD4low precursor population or, alternatively, that both
CD8+ and CD8 DCs derive from an
independent nonlymphoid, nonmyeloid DC precursor. In conclusion,
although we favor the hypothesis that both CD8+ and
CD8 DCs derive from a lymphoid-committed precursor, a
precise study of the differentiation process of CD8+ and
CD8 DCs is required to define conclusively their origin.
(Blood. 2000;96:2511-2519)
© 2000 by The American Society of Hematology.
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Introduction |
Two main dendritic cell (DC) subsets have been
described in the mouse, which can be distinguished on the basis of
their differential CD8 expression,1,2 The finding that
murine thymic DCs expressing CD8 derived from CD4low
lymphoid precursors, devoid of myeloid differentiation
potential,3 lead to the definition of the lymphoid
DC lineage.
Over the past few years, numerous reports dealing with the phenotype,
localization, and function of murine CD8+ DCs and
CD8 DCs have been published, contributing enormously
to our understanding of DC biology.4-15 However, certain
issues dealing with CD8+ versus CD8
DCs, such as their differential T-cell stimulatory capacity, production
of cytokines, responsiveness to chemokines, induction of TH1 versus TH2
responses, and, in particular, the definition of their respective
precursor populations, are still controversial or remain unknown.
These studies were based on a variety of experimental approaches
performed both in vivo and in vitro and were undertaken by using
different DC purification techniques. Consequently, some published
controversial results regarding the phenotype and/or the function of
the different DC subsets might reflect differences in the DC isolation
method used. The protocols of DC isolation used by different research
groups differ mainly in the enzymatic digestion performed, in the way
of obtaining a DC-enriched low-density fraction, and, more importantly,
in the antibodies used to eliminate contaminating cells by magnetic or
flow cytometry separation procedures. This negative selection of
contaminating cells is of special relevance because certain DC
populations can be completely or partially lost, depending on the
antibodies used. Therefore, the design of an appropriate negative
selection protocol requires an exhaustive phenotypic study of the DC
subset to be purified. In this sense, the majority of the reports
dealing with the function of CD8+ versus
CD8 DC subsets4-7,9 and, more importantly,
the article establishing that CD8+ DCs but not
CD8 DCs derived from lymphoid committed
precursors,16 which has been the basis of the concept of
murine lymphoid versus myeloid DCs, rely on a DC isolation method using
anti-CD4 and anti-F4/80 antibodies to deplete CD4+ T cells
and F4/80+ macrophages. In the present report, we have
performed a precise phenotypic analysis of murine splenic
CD8+ and CD8 DCs with regard to the
expression of certain conflictive markers, and we have re-addressed the
question of the derivation of both DC subsets from CD4low
precursors. Our results revealed that CD8 DCs, but not
CD8+ DCs, expressed CD4 and F4/80 and, therefore, that
the use of antibodies against those molecules during
CD8 DC isolation might lead to the loss of a
significant proportion of them. More importantly, CD8
DCs together with CD8+ DCs were generated from
lymphoid-committed CD4low precursors in irradiation
chimeras in which the development of donor-type DCs was analyzed on
DC-enriched splenic low-density fractions, without depleting cells
expressing CD4 and/or F4/80. In irradiation chimeras donor-type
CD8+ and CD8 DCs displayed an
equivalent phenotype to that of their control counterparts and were
generated from CD4low precursors at a similar
CD8+-to-CD8 ratio to that found in control
mice. These results suggest that both CD8+ and
CD8 DCs derive from a common precursor and do not
support the concept in the murine system of the CD8+
lymphoid-derived and CD8 myeloid-derived DC lineages.
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Materials and methods |
Mice
For phenotypic and functional experiments 5- to 6-week-old C57
BL/6 and BALB/c mice were used. In reconstitution experiments, donors
were 5- to 6-week-old C57 BL/Ka Ly 5.2 mice and recipients were
8-week-old C57 BL/6 Ly 5.1 Pep3b mice.
Preparation of DC-enriched very low density cell fractions
Spleens were cut into small fragments and digested with
collagenase A (0.5 mg/mL; Boehringer-Mannheim, Mannheim, Germany) and
Dnase I (40 µg/mL; Boehringer-Mannheim) in RPMI 1640 medium supplemented with 5% fetal calf serum (FCS) for 10 minutes at 37°C
with continuous agitation. Digested fragments were filtered through a
stainless-steel sieve, and cell suspensions were washed twice in
phosphate-buffered saline (PBS) supplemented with 5% FCS and 5 mmol/L
EDTA (PBS-EDTA-FCS) containing 5 µg/mL Dnase I. The cells were
resuspended in cold isoosmotic Optiprep solution (Nyegaard Diagnostics,
Oslo, Norway), pH7.2, density 1.061 g/cm3, containing 5 mmol/L EDTA to dissociate DC-thymocyte complexes, and a very
low-density cell fraction (1.061-density fraction), accounting for less
than 1% of the starting cell population, obtained by centrifugation at
1700g for 10 minutes, and washed twice in PBS-EDTA-FCS.
Preparation of CD8 DC-enriched cell
fractions
CD8 DC-enriched cell fractions were obtained by
treating DC-enriched 1.061-density fractions for 50 minutes at 4°C
with a monoclonal antibody (mAb) mixture, including anti-CD3 (clone
KT3-1.1), anti-CD8 (clone 53-6.72), anti-B220 (clone RA3-6B2), and
anti-granulocyte antigen Gr1 (clone RB6-8C5). The unwanted cells were
removed magnetically after incubation for 30 minutes at 4°C with
anti-rat immunoglobulin-coated magnetic beads (Dynabeads, Dynal, Oslo,
Norway) at a 7:1 bead-to-cell ratio. Analysis of CD11c versus CD8
expression of CD8 DC-enriched cell fractions revealed
that they were composed of more than 80% CD11c+
CD8 DCs and approximately 20% CD11c
CD8 contaminants, CD11c+
CD8+ DCs, representing less than 1% (data not shown).
In vivo depletion with anti-CD4 antibodies
Depletion of CD4+ cells was achieved by
intraperitoneal injection of the in vivo depleting anti-CD4 antibody
GK1.5. For this purpose, mice received three injections of 300 µg of
GK1.5 on 3 consecutive days and were analyzed 24 hours after the
last injection.
Modulation of cell surface marker expression by
CD8 DCs on culture
CD8 DC-enriched cell fractions prepared as
described above were cultured for 24 hours or 48 hours at 37°C in the
presence 100 µg/mL anti-CD40 mAb (clone FGK45) or anti-CD43 (clone
S7). The culture medium was RPMI 1640 supplemented with 10% FCS, 10 mmol/L Hepes, 50 µmol/L 2-mercaptoethanol, 100 U/mL
penicillin-streptomycin, and 100 ng/mL granulocyte-macrophage
colony-stimulating factor (GM-CSF).
MLR assay
Splenic CD4 CD8 DCs,
CD4+ CD8 DCs, total CD8
DCs, or CD8+ DCs from C57BL/6 (H-2b) mice
were cultured with purified T cells obtained from mesenteric lymph
nodes of BALB/c (H-2d) mice in flat-bottom 96-well plates
(1 × 105 cells per well) at different APC-to-T cell
ratios. T-cell proliferation was assessed after 5 days by
[3H] thymidine (1 µCi/well) uptake in a 4-hour pulse or
by CD25 expression. For this assay, CD4
CD8 DCs, CD4+ CD8 DCs,
and total CD8 DCs were sorted from
CD8 DC-enriched cell fractions. CD8+
DCs were sorted from DC-enriched 1.061-density fractions. The sorted
populations had a purity of more than 97%.
Isolation of CD4low and CD44+
CD25+ precursor populations
CD4low precursors were isolated from C57 BL/Ka Ly
5.2 donor thymuses by depleting pre-T cells, double-positive and
single-positive thymocytes, B cells, DCs, macrophages, and granulocytes
by complement-mediated cytotoxicity by using anti-CD3 (clone Y-CD3) and
anti-CD8 (clone 31M) and then immunomagnetic bead depletion after
incubation with anti-CD3 (clone KT3), anti-CD8 (clone 53.6.7-2),
anti-CD25 (clone PC61.5), anti-B220 (clone RA3-6B2), anti-MHC class II
(MHC II) (clone FD11), anti-macrophage antigen F4/80 (clone C1.A3.1),
and anti-granulocyte antigen Gr-1 (clone RB6-8C5). CD4low
precursors were then sorted as Thy-1low CD44+
cells after double immunofluorescent staining fluorescein
isothiocyanate (FITC)-conjugated anti-Thy-1 (clone AT15) and
phycoerythrin (PE)-conjugated anti-CD44 (clone IM7, Pharmingen, San
Diego, CA). Flow cytometric cell sorting was carried out on a FACSort
instrument (Becton Dickinson, Mountain View, CA). The sorted
preparation had a purity of more than 98% and contained less than 1%
CD11c+ cells as assessed after staining with biotinylated
anti-CD11c (clone N418; data not shown) followed by
streptavidin-tricolor (Caltag, San Francisco, CA).
CD44+ CD25+ precursors were isolated by
complement-mediated cytotoxicity by using anti-CD3 (clone Y-CD3),
anti-CD4 (clone 172.4), and anti-CD8 (clone 31M) and then sorting after
double immunofluorescent staining with PE-conjugated anti-CD44 and
biotinylated anti-CD25 followed by streptavidin-tricolor. The sorted
population had a purity of more than 97%.
Reconstitution experiments with CD4low precursors or
CD44+ CD25+ precursors
Thymic CD4low precursors (3 × 104 )
or thymic CD44+ CD25+ precursors
(3 × 104) from C57 BL/Ka Ly 5.2 donor mice were injected
intravenously into  -irradiated (7 Gy) C57 BL/6 Ly 5.1 Pep3b recipient mice, along with 3 × 104 Ly
5.1 bone marrow (BM) cells to ensure survival of recipients.
Reconstitution experiments with BM cells
BM cells (2 × 106) from C57 BL/Ka Ly 5.2 donor
mice were injected intravenously into -irradiated (7 Gy) C57 BL/6 Ly
5.1 Pep3b mice.
Flow cytometry
DC-enriched very low density cell fractions (Figure
1) were analyzed after triple staining
with FITC-conjugated anti-CD11c (clone N418), PE-conjugated anti-CD8
(clone CT-CD8a, Caltag), and biotin-conjugated anti-DEC-205 (clone
NLDC-145); anti-macrophage antigen F4/80 (clone 31-A3-1); or anti-CD4
(clone GK1.5) followed by streptavidin-tricolor (Caltag). The
phenotypic analysis of CD4 CD8 DCs and
CD4+ CD8 DC subsets (Figure
2) was performed after triple staining
with FITC-conjugated anti-CD11c, PE-conjugated anti-CD4 (clone CT-CD4, Caltag), and biotin-conjugated anti-DEC-205; anti-macrophage antigen F4/80; anti-Mac-1 (clone M1/70); anti-FcRII/III (clone 2-4G2); anti-LFA-1 (clone FD441.8); anti-CD69 (clone H.1.2F3); anti-B7-2 (clone GL1, Pharmingen); anti-CD40 (clone FGK45); or anti-MHC class II
(clone FD11-54.3), followed by streptavidin-tricolor. Analysis of cell
surface marker expression by CD8 DCs on culture
(Figure 3) was performed after triple
staining with FITC-conjugated anti-CD11c, PE-conjugated anti-CD8,
and biotin-conjugated anti-CD4; anti-macrophage antigen F4/80; or anti-DEC-205 followed by streptavidin-tricolor. Analysis of T-cell proliferation (Figure 4) was performed
after triple staining with FITC-conjugated anti-CD8, PE-conjugated
anti-CD4, and biotin-conjugated anti-CD25 (clone PC61.5) followed by
streptavidin-tricolor. Analysis of DC reconstitution (Figures
5 and 6)
was performed on splenic DC-enriched 1.061-density
fractions after triple staining with FITC-conjugated anti-CD11c,
PE-conjugated anti-Ly 5.2 (clone AL1-4A2, Pharmingen), and
tricolor-conjugated anti-CD8 (clone CT-CD8a, Caltag). Because, after
transfer of BM precursors, more than 95% of DCs were of donor origin,
the phenotypic analysis of Ly 5.2+ CD8 DCs
(Figure 6) was performed after triple staining with
FITC-conjugated anti-CD11c, PE-conjugated anti-CD8, and
tricolor-conjugated anti-CD4 (clone CT-CD4, Caltag) or
biotin-conjugated anti-macrophage antigen F4/80 followed by
streptavidin-tricolor. Analysis was performed on a FACSort flow
cytometer.
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| Figure 1.
CD4 and F4/80 expression by splenic DCs.
(A) Phenotype of CD8+ and CD8 DCs
analyzed on splenic DC-enriched very low density fractions obtained
without antibody-mediated depletion. Black histograms show the
proportion of CD8 and CD8+ DCs after
gating for CD11c+ cells and the proportion of
CD4 and CD4+ DCs within the
CD8 subset. White profiles represent control
stainings. The percentage of cells positive for the indicated markers
is shown under the gray histograms. (B) Splenic CD11c versus CD8
profiles of control mice and mice injected intravenously with in vivo
depleting anti-CD4 antibodies, showing that most CD8
DCs were eliminated after anti-CD4 treatment. Data are representative
of 3 experiments with similar results.
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| Figure 2.
Comparative phenotypic analysis of CD4
CD8 versus CD4+ CD8 DCs.
The phenotype of CD4 CD8 versus
CD4+ CD8 DCs was performed on
CD8 DC-enriched fractions after gating for
CD11c+ CD4 or CD11c+
CD4+ cells. The percentage of cells positive for the
indicated markers is shown. In addition, the percentage of
CD4 CD8 DCs expressing high levels of
FcR or LFA-1 is also indicated.
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| Figure 3.
Modulation of CD4 expression by CD8 DCs
on culture.
CD8 DC-enriched fractions were cultured for 24 hours
or 48 hours alone or in the presence of anti-CD40 or anti-CD43
antibodies. CD8 DCs were analyzed after gating for
CD11c+ CD8 cells. The percentage of cells
positive for the indicated markers as well as the mean fluorescence
intensity for CD11c expression is shown. White profiles represent
control stainings. Data are representative of 3 experiments with
similar results.
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| Figure 4.
T-cell stimulation capacity of CD8+ DCs
versus CD8 DCs.
T-cell stimulatory capacity of CD8+ versus
CD8 DC (A) and CD4 CD8
versus CD4+ CD8 DC (B). DCs from C57BL/6
mice were cultured with purified allogeneic T cells from BALB/c mice at
the indicated APC:T cell ratios. After 5 days, T-cell proliferation was
determined by [3H] thymidine uptake or CD25 expression.
Data are representative of 3 experiments with similar results. In the
[3H] thymidine incorporation assay, error bars represent
the SD for triplicate cultures.
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| Figure 5.
Reconstitution of splenic CD8+ and CD8
DCs from CD4low lymphoid precursors.
Thymic CD4low precursors (3 × 104) from C57
BL/Ka Ly 5.2 donor mice were injected intravenously into -irradiated
(7 Gy) C57 BL/6 Ly 5.1 Pep3b recipient mice, along with
4 × 104 Ly 5.1 BM cells to ensure survival of
recipients. At the indicated times, mice were analyzed for
donor-derived DCs, identified as Ly 5.2+ CD11c+
cells, in splenic DC-enriched 1.061-density fractions. The percentage
of Ly5.2 and Ly5.2+ CD11c+ DCs
(contour plots), as well as the percentage of CD8 and
CD8+ DCs among Ly5.2+ DCs (black histograms)
are indicated. Grey profiles show the expression of CD4 and F4/80 by
CD8 DCs. White profiles represent control stainings.
These results are representative of four independent experiments with
similar results.
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| Figure 6.
Reconstitution of splenic CD8+ and
CD8 DCs from CD44+ CD25+
precursors.
Thymic CD44+ CD25+ precursors
(3 × 104) were injected intravenously into 7 Gy
-irradiated 8-week-old C57 BL/6 Ly 5.1 Pep3b recipient
mice, along with 4 × 104 Ly 5.1 BM cells to ensure
survival of recipients. Ten days after transfer of precursors, mice
were analyzed for donor-derived DCs. The percentage of
Ly5.2+ CD11c+ DCs as well as the percentage of
CD8 and CD8+ DCs among
Ly5.2+ DCs are indicated. Data are representative of 2 independent experiments with similar results.
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Results |
Phenotypic profile of mouse splenic DC revisited: CD4 and F4/80
expression by CD8 splenic DCs
The most comprehensive analyses of mouse DC phenotype reported so
far1,2 have been performed on highly enriched DC
preparations obtained after immunomagnetic depletion of contaminating
cells with specific mAbs. Those contaminants comprise essentially T and
B lymphocytes, macrophages, and granulocytes, and, therefore, mAbs used
for depleting them included usually at least anti-CD3, anti-CD4,
anti-B220, anti-FcR, anti-F4/80 (macrophage marker), and anti-Gr-1
(granulocyte marker). Obviously, the selection of these mAbs was
originally made on the basis of the assumption that DCs did not express
those cell surface molecules.
However, DCs can be accurately analyzed on DC-enriched very low-density
cell fractions, obtained without including the mAb-mediated magnetic
bead depletion step, by using a centrifugation medium adjusted at 1.061 g/mL. As shown in Figure 1A, the phenotypic analysis of splenic DCs
performed under those conditions revealed that CD8 DCs
and CD8+ DCs represented approximately 70% and 30% of
all splenic DCs, respectively. As shown in the
Table, estimation of the absolute DC
number revealed that the spleen of 5- to 6-week-old C57 BL/6 mice
contained 180 000-190 000 CD8 DCs and
80 000-90 000 CD8+ DCs. Interestingly,
CD8 DCs expressed the macrophage marker F4/80 and more
interestingly 70%-75% of them were CD4+. Moreover,
CD8+ DCs were essentially negative for F4/80 and CD4,
although approximately 20% of them expressed low levels of these
markers. Consequently, in the mouse spleen CD8+, DCs
were CD4 F4/80 DEC-205+,
whereas CD8 DCs were CD4+
F4/80+ DEC-205. However, within the lymph
nodes, an additional DC subset expressing intermediate levels of CD8
exists, which has been related to Langerhans cells.2 These
cells displayed the same phenotype as splenic CD8+ DCs
regarding the markers considered above, that is, they were CD4 F4/80 DEC-205+ (data not shown).
To check whether CD4 was functionally expressed at the surface of
splenic CD8 DCs, BALB/c mice were injected
intraperitoneally with the in vivo depleting anti-CD4 antibody GK1.5.
As shown in Figure 1B, the spleen of mice treated in vivo with GK1.5
suffered a profound depletion of CD8 DCs, whereas the
CD8+ DC subset was unaffected. CD8 DCs
remaining after GK1.5 injection most likely corresponded to
CD8 DCs negative for CD4 or expressing low levels of
this molecule. This result indicates that the CD4 molecule was
functionally expressed by splenic CD8 DCs.
In conclusion, the phenotypic analysis of splenic DCs performed on
DC-enriched very low-density cell fractions revealed that expression of
CD8 and CD4 by splenic DC subsets is mutually exclusive, in that
CD8+ DCs are CD4 and, inversely,
CD8 DCs are CD4+.
Comparative phenotypic analysis of CD4
CD8 DCs versus CD4+
CD8 DCs
To investigate whether the CD4 and CD4+
cells CD8 DC corresponded to distinct DC populations
or to different CD4 expression levels within the CD8
DC subset, we performed a comparative phenotypic study of
CD4 CD8 DCs versus CD4+
CD8 DCs. For this purpose, a CD8
DC-enriched fraction was obtained from a splenic 1.061-density fraction
after immunomagnetic bead depletion with anti-CD3, anti-CD8, anti-B220, and anti-Gr-1 mAbs. The analysis was performed on this CD8 DC-enriched fraction by gating on
CD4 or CD4+ cells (as shown in Figure 1A)
after triple immunofluorescence staining with FITC-conjugated
anti-CD11c, PE-conjugated anti-CD4, and biotin-conjugated antibodies
against the cell surface markers indicated in Figure 2. Our data show
that CD4 CD8 and CD4+
CD8 DCs have a very similar phenotypic profile with
regard to a variety of cell surface molecules, including DC and
macrophage markers, adhesion, activation, and costimulatory molecules.
There was, however, a slight but significant difference with regard to
the expression of FcR (CD16-CD32) and LFA-1, because approximately 30%
CD4 CD8 DCs but not CD4+
CD8 DCs expressed high levels of these markers.
Modulation of CD4 expression by CD8 DCs
on culture
The data derived from the comparative phenotypic analysis of
CD4 versus CD4+ CD8 DC
subsets suggest that they belong to a unique DC category with differential expression of CD4. Therefore, it can be speculated that
CD4 expression levels correlate with different activation and/or
maturation states. To test this hypothesis, CD8
DC-enriched populations were cultured alone or in the presence of
antibodies against the molecules CD40 or CD43, known to induce DC
activation on ligation.17-19 As illustrated in Figure 3,
CD4 expression was strongly down-regulated in CD8 DCs
after 24 hours in culture and was almost undetectable after 48 hours.
Moreover, addition of anti-CD40 or anti-CD43 antibodies did not prevent
CD4 down-regulation, suggesting that CD4 expression by
CD8 DCs was not related to the activation of these
cells. Under the same experimental conditions, F4/80 expression was
also down-regulated after 48 hours, although a significant proportion
of CD8 DCs remained positive for this marker. However,
CD11c underwent only a slight down-regulation on culture. As for CD4,
anti-CD40 or anti-CD43 antibodies had no effect on F4/80 or CD11c
expression on 48-hour culture.
T-cell stimulation capacity of CD8+ versus
CD8 DCs
To test whether CD4 expression by CD8 DCs was
correlated with their functional potential as antibody-presenting
cells, we tested their capacity to induce T-cell stimulation in an MLR
assay. For these experiments, CD4 and CD4+
subsets of CD8 DCs, as well as total
CD8 DCs and CD8+ DCs, were FACS-sorted
and cultured with purified allogeneic T cells. After 5 days, T-cell
proliferation was determined by [3H] thymidine uptake or
by CD25 expression. We first tested the differential T-cell stimulatory
potential of CD8 versus CD8+ DCs. Our
data revealed that CD8+ DCs induced a higher
[3H] thymidine uptake and CD25 up-regulation in an
allogeneic MLR than CD8 DCs (Figure 4A). Because, as
shown above, there was a direct correlation between [3H]
thymidine incorporation and expression of CD25, the latter was
subsequently used to assess, in the same in vitro assay, the capacity
of CD4 CD8 DCs versus CD4+
CD8 DCs to stimulate CD8+ or
CD4+ T cells. As illustrated in Figure 4B,
CD4 and CD4+ CD8 DCs
displayed a similar T-cell stimulatory potential in MLR, although the
CD4+ CD8 DC subset induced a slightly
higher response of both CD8+ and CD4+ T cells.
Interestingly, preliminary results indicate that the endocytic capacity
of CD8 DCs appears to be restricted to the
CD4+ subset (data not shown).
Reconstitution of splenic CD8+ DCs and
CD8 DCs from CD4low lymphoid
precursors
As discussed above, in the murine system the concept that
CD8 DCs and CD8+ DCs represent the
myeloid and lymphoid DC subsets, respectively, derives essentially from
a report analyzing the DC reconstitution potential of
CD4low precursors on intravenous injection.16
In that report, only CD8+ DCs were found among the
progeny of CD4low precursor after 2 weeks. However, in this
study DC reconstitution was analyzed after DC purification, using an
immunomagnetic bead depletion protocol employing anti-CD4 and
anti-F4/80 antibodies, and, therefore, CD8 DCs could
have been excluded from the analysis.
To test this hypothesis and thus to investigate whether
CD8 DCs derive from the lymphoid lineage-committed
CD4low precursor population, CD4low precursors
isolated from Ly 5.2 mice were transferred intravenously into
-irradiated Ly 5.1 recipient mice. These mice were subsequently analyzed for donor-derived DCs, identified as Ly 5.2+
CD11c+ cells, in DC-enriched 1.061-density fractions,
obtained without including the mAb-mediated magnetic bead depletion
step, as described above (Figure 1A). Our data, shown in Figure 5,
demonstrate that 7 days after the transfer approximately 70% of DCs
were donor derived and, interestingly, that among those cells both
CD8+ and CD8 DCs were found. At this
time point, Ly 5.2+ CD8+ and Ly
5.2+ CD8 DCs represented 55% and 45% of
all Ly 5.2+ DCs, respectively. Importantly, donor-derived
CD8 DCs derived from CD4low precursors
displayed the phenotypic profile characteristic of CD8
DCs found in control spleen (Figure 1), that is, they expressed high
levels of CD4 and F4/80. After 14 days, only 30% Ly 5.2+
DCs were found, indicating that the DC reconstitution potential of the
CD4low precursor population was partially extinguished and
replaced by that of Ly 5.1+ BM precursors. Both Ly
5.2+ CD8+ and Ly 5.2+
CD8 DCs were also found after 14 days, each subset
constituting approximately 50% of all Ly 5.2+ DCs.
Therefore, as postulated above, in the report by Wu et
al,16 CD8 DCs of donor origin could have
been missed in the analysis because of their CD4 and/or F4/80
expression that could have determined their depletion, because
antibodies against these molecules were used for immunomagnetic
depletion in this study. In conclusion, these data indicate that both
CD8 and CD8+ DCs are generated from
CD4low lymphoid-committed precursors. To exclude that
CD8 DCs could have been transferred with the
CD4low precursors, the CD4low precursor
population was analyzed before transfer for the presence of
CD11c+ cells and no CD11c+ cells were found
(data not shown). In addition, mice reconstituted with
CD4low precursors were analyzed 2 days after transfer for
Ly 5.2+ DCs, but no CD11c+ were found in the
spleen at this time point (data not shown). Therefore, these data
reinforce the view that Ly 5.2+ CD8 and
CD8+ DCs derived from the transferred CD4low precursors.
To further strengthen our data with CD4low precursors, we
tested the ability of the next downstream precursor population, namely the CD44+ CD25+ pro-T cell precursors that have
lost the capacity to form B cells but still form DCs16 to
generate CD8 and CD8+ DCs on
intravenous transfer. As shown in Figure 6, 10 days after the transfer
of CD44+ CD25+ precursors, approximately 30%
of CD11c+ DCs were donor derived, and both
CD8+ and CD8 DCs were found.
Interestingly, CD8+ and CD8 DCs
represented 58% and 42% of Ly 5.2+ DCs, respectively,
indicating that CD4low and CD44+
CD25+ precursors generated the two splenic DC subsets at
comparable CD8 DC-to-CD8+ DC ratios.
Kinetics of splenic DC differentiation and phenotypic variations of
CD8 DCs during reconstitution after transfer of
BM cells
To study the phenotype of CD8 DCs during the
process of DC reconstitution after irradiation, BM cells from Ly 5.2 donor mice were transferred intravenously into -irradiated Ly 5.1 recipient mice. These mice were subsequently analyzed for donor-derived DCs at 7, 14, and 21 days after transfer. For this purpose, the experiments were carried out with BM precursor cells instead of CD4low precursors because the progressive loss of DC
reconstitution capacity of the latter, as a result of their extinction,
does not allow the analysis of the phenotypic variations undergone by
CD8 DCs during the reconstitution process.
As shown in Figure 7, during the
reconstitution period analyzed (from 7 to 21 days after transfer),
virtually all the DCs present in the spleen of reconstituted mice were
of donor origin; only after 21 days a small proportion of Ly
5.1+ CD11c+ cells (approximately 5% of all
CD11c+ cells) was detected. With regard to the proportion
of the CD8 and CD8+ subsets, at 7 and
14 days CD8+ DCs represented approximately 50% of
splenic DCs, but after 21 days the proportion of CD8
DCs increased, with the CD8 DC-to-CD8+
DC ratio being 70:30 (ie, as in control spleens; Table 1).
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| Figure 7.
Kinetics of splenic DC differentiation and phenotypic
variations of CD8 DCs during reconstitution after
transfer of BM cells.
BM cells (2 × 106) from C57 BL/Ka Ly 5.2 donor mice were
injected intravenously into -irradiated (7 Gy) C57 BL/6 Ly 5.1 Pep3b recipient mice. At the indicated times, mice were
analyzed for donor-derived DCs in splenic DC-enriched 1.061-density
fractions. The percentage of Ly5.2 and Ly5.2+
CD11c+ DCs (contour plots), as well as the percentage of
CD8 and CD8+ DCs among
Ly5.2+ DCs (black histograms) are indicated. Grey profiles
show the expression of CD4 and F4/80 by Ly5.2+
CD8 DCs; the percentage of CD4+ or
F4/80+ cells is indicated. White profiles represent control
stainings. These results are representative of three independent
experiments with similar results.
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With regard to kinetics of reconstitution of the two DC subsets after
BM transfer, the estimation of the absolute numbers of
CD8 and CD8+ DCs of donor origin
revealed that 7 days after transfer both DC compartments were fully
reconstituted, with the absolute number of CD8+ DCs
being 2-fold higher than in control spleens (Table 1). The absolute
number of both DC subpopulations increased considerably during the
second week of reconstitution, and after 3 weeks, even though the
CD8 DC-to-CD8+ DC ratio was similar to
that of control spleens, the absolute number of both DC subsets was
still 2-fold to 3-fold higher than in controls.
Globally, the data derived from our experiments of reconstitution after
irradiation show that CD8 and CD8+ DC
reconstitution is faster than that of T and B lymphocytes, that the
process of DC reconstitution occurs simultaneously for CD8 and CD8+ DCs, and that both subsets
are produced in equal numbers during the first 2 weeks after transfer
of precursors. In addition, the data reveal that CD8
and CD8+ DCs are generated at the same ratio during
reconstitution with both CD4low precursors and BM precursors.
Concerning the phenotype of reconstituted DCs, from the shortest time
point analyzed, CD8+ DCs displayed the same phenotypic
profile as their control counterparts (not shown). However, as
illustrated in Figure 7, CD8 DCs underwent variations
in their CD4 and F4/80 expression during reconstitution because the
percentage of CD4+ CD8 |
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Abstract
Introduction